Emerging Infectious Diseases [Volume 5 No.1 / January - February 1999] The January–February 1999 issue of the journal will contain the articles listed below. Highlighted articles are currently available in electronic format. To view a 63K image of the full cover click here. International Update * Emerging Viral Diseases: An Australian Perspective, John S. Mackenzie Perspectives * The Economic Impact of Staphylococcus aureus Infection in New York City Hospitals, R. J. Rubin, C. A. Harrington, A. Poon, K. Dietrich, J. A. Greene, and A. Moiduddin * Socioeconomic and Behavioral Factors Leading to Acquired Bacterial Resistance to Antibiotics in Developing Countries, Iruka N. Okeke, Adebayo Lamikanra, and Robert Edelman * Campylobacter jejuni—An Emerging Foodborne Pathogen, Sean F. Altekruse, Norman J. Stern, Patricia I. Fields, and David L. Swerdlow Synopses * Comparative Genomics and Host Resistance Against Infectious Diseases, Salman T. Qureshi, Emil Skamene, and Danielle Malo * Cyclospora: An Enigma Worth Unraveling, Charles R. Sterling and Ynés R. Ortega * Using Monoclonal Antibodies to Prevent Mucosal Transmission of Epidemic Infectious Diseases, Larry Zeitlin, Richard A. Cone, and Kevin J. Whaley Research * HIV-1 Dual Infections and Recombinants Are an Integral Part of the HIV Epidemic in Brazil, Artur Ramos, Amilcar Tanuri, Mauro Schechter, Mark A. Rayfield, Dale J. Hu, Maulori C. Cabral, Claudiu I. Bandea, James Baggs, and Danuta Pieniazek * Genetic Diversity and Distribution of Peromyscus-Borne Hantaviruses in North America, Martha C. Monroe, Sergey P. Morzunov, Angela M. Johnson, Michael D. Bowen, Harvey Artsob, Terry Yates, C.J. Peters, Pierre E. Rollin, Thomas G. Ksiazek, and Stuart T. Nichol * Climatic and Environmental Patterns Associated with Hantavirus Pulmonary Syndrome Cases in the Four Corners Region, David M. Engelthaler, David G. Mosley, James E. Cheek, Craig E. Levy, Kenneth K. Komatsu, Paul Ettestad, Ted Davis, Dale T. Tanda, Lisa Miller, J. Wyatt Frampton, Richard Porter, and Ralph T. Bryan The January-March 1999 issue of Emerging Infectious Diseases will be highlighting hantavirus research. This theme issue will include the following cluster of articles: * Long-Term Studies of Hantavirus Reservoir Population in the Southwestern United States: Rationale, Potential, and Methods, James N. Mills,Thomas G. Ksiazek, C.J. Peters, and James E. Childs * Long-Term Hantavirus Persistence in Rodent Populations in Central Arizona, Ken D. Abbott, Thomas G. Ksiazek, and James N. Mills * A Longitudinal Study of Sin Nombre Virus Prevalence in Rodents, Southeastern Arizona, Amy J. Kuenzi, Michael L. Morrison, Don E. Swann, Paul C. Hardy, and Giselle T. Downard * Long-Term field Studies of Small Mammals for Hantavirus Prevalence: Statistical Sensitivity for Detection of Spatial and Temporal Patterns in Rodent Population Densities, Cheryl A. Parmenter, Terry L. Yates, Robert R. Parmenter, and Jonathan L. Dunnum * Natural History of Sin Nombre Virus in Western Colorado, Charles H. Calisher, William Sweeney, James N. Mills, and Barry J. Beaty * Long-Term Studies of Hantavirus Reservoir Populations in the Southwestern United States: A Synthesis, James N. Mills, Thomas G. Ksiazek, C.J. Peters, and James E. Childs Dispatches * Proficiency of Clinical Laboratories in and Near Monterrey, Mexico to Detect Vancomycin-Resistant Enterococci, L. Clifford McDonald, Luis R. Garza, and William R. Jarvis * Staphylococcus aureus with Reduced Susceptibility to Vancomycin Isolated from a Patient with Fatal Bacteremia, Sharon S. Rotun, Virginia McMath, Dianna J. Schoonmaker, Peggy S. Maupin, Fred C.Tenover, Bertha C. Hill, and David A. Ackman * Candida dubliniensis Candidemia in Patients with Chemotherapy-Induced Neutropenia and Bone Marrow Transplantation, Jacques F.G.M. Meis, Markus Ruhnke, Ben E. De Pauw, Frank C. Odds, Wolfgang Siegert, and Paul E. Verweij * Household Transmission of Streptococcus pneumoniae, Alberta, Canada, James D. Kellner, A. Patrick Gibb, Jenny Zhang, and Harvey R. Rabin * Preventing Zoonotic Diseases in Immunocompromised Persons: The Role of Physicians and Veterinarians, Sara Grant and Christopher W. Olsen * Mycoplasma penetrans Bacteremia Associated with Primary Antiphospholipid Syndrome, Antonio Yáñez, Lilia Cedillo, Olivier Neyrolles, Encarnación Alonso, Marie-Christine Prévost, Jorge Rojas, Harold L. Watson, Alain Blanchard, and Gail H. Cassell * Infectious Diarrhea in Tourists Staying in a Resort Hotel , Rachel M. Hardie, Patrick G. Wall, Patricia Gott, Madhu Bardhan, and Christopher L.R. Bartlett Commentary * Hantavirus Pulmonary Syndrome—A Mid-Course Assessment, Robert E. Shope Letters * Navigational Instinct: A Reason Not to Live Trap Deer Mice in Residences, Charles H. Calisher, William P. Sweeney, J. Jeffrey Root, and Barry J. Beaty * Bartonella quintana in Body Lice Collected from Homeless Persons in Russia, Elena B. Rydkina, Véronique Roux, Eugenia M. Gagua, Alexandre B. Predtechenski, Irina V. Tarasevich, and Didier Raoult * Tick-Transmitted Infections in Transvaal: Consider Rickettsia africae, Pierre-Edouard Fournier, Jean Beytout, and Didier Raoult * Extended-Spectrum Beta-Lactamase-Producing Salmonella Enteritidis in Trinidad and Tobago, B.P. Cherian, Nicole Singh, W. Charles, and P. Prabhakar * New emm (M Protein Gene) Sequences of Group A Streptococci Isolated from Malaysian Patients, Farida Jamal, Sabiha Pit, Richard Facklam, and Bernard Beall * Mutant Chemokine Receptor (CCR-5) and its Relevance to HIV Infection in Arabs, Iman H. Al-Sheikh, Amjad Rahi, and Mohammed Al-Khalifa News and Notes * Workshop on the Potential Role of Infectious Agents in Cardiovascular Disease and Atherosclerosis * Workshop on Risks Associated with Transmissible Spongiform Encephalopathies (TSEs) * USDA-Report on Potential for International Travelers to Transmit Foreign Animal Diseases to U.S. Livestock or Poultry * Second Annual Conference on Vaccine Research, March 1999 * International Scientific Forum on Home Hygiene ——————————————————————————————————————————————————————————————————————————— Commentary ——————————————————————————————————————————————————————————————————————————— Update International Editors --------------------------------------------------------------------------- Emerging Viral Diseases: An Australian Perspective John S. Mackenzie The University of Queensland, Brisbane, Australia [Fig] Figure. Dr. Mackenzie is professor and head of the Department of Microbiology and Parasitology, University of Queensland, Brisbane, Australia. His research interests include the epidemiology, ecology, and molecular biology of mosquito-borne and emerging zoonotic viruses. With a few exceptions, emerging diseases in Australia are similar to those in other industrialized countries (1-8). Most exceptions are either vector-borne or zoonotic viral diseases, the major focus of this update. The continuing emergence of antibiotic resistance is a worldwide problem. In Australia, antibiotic resistance is being reported from a growing number of organisms (9-15), often necessitating new case management practices and guidelines (16,17). Also, like other countries, Australia has had a number of foodborne (18-22) and waterborne (23-25) epidemics in the past few years; the major difference is that the higher incidence of enterohemorrhagic Escherichia coli linked to outbreaks of hemolytic uremic syndrome is associated with serotype O111:H- rather than serotype O157:H-, which is more common in other countries. Major waterborne epidemics or contamination of reservoirs due to Cryptosporidium parvum have occurred over the past 3 years in the Eastern States of Australia (23-25), with the largest and most recent being a problem of contamination (in association with Giardia lamblia) in the Sydney water supply between July and September, 1998. However, despite this contamination, no increases in the number of cases of diarrheal disease were reported, perhaps because the inhabitants of Sydney were advised to boil the water before drinking it (25). The distribution and incidence of most of the recently described viral diseases (e.g., human herpesviruses 6–8 and hepatitis C and E viruses) in Australia are similar to those reported in other industrialized nations (3). Recent data for hepatitis G in selected Australian populations also support this contention (26). Vector-Borne Viral Diseases Australia has more than 70 arboviruses, but relatively few cause human disease (27,28). The most common arbovirus causing human disease is Ross River virus, an alphavirus, which causes an epidemic polyarthritis. Although Ross River virus incidence has increased over the past decade, the virus is not emerging; its increased incidence is probably due to increased awareness and recognition by general practitioners, improved diagnostic reagents, and increasing encroachment of human habitation into or near wetlands and other areas conducive to mosquito breeding. The only indigenous virus that can be called "emerging" is Barmah Forest virus, also an alphavirus and also the cause of an epidemic polyarthritis-like disease. Associated with human disease only since 1988 and increasing in incidence as diagnostic reagents have become available and clinicians have become aware of it, the virus has spread into new geographic areas, such as Western Australia (29,30). The two mosquito-borne diseases of particular concern, however, are "imports"—Japanese encephalitis (JE) and dengue viruses. Japanese Encephalitis Virus The first outbreak of JE in the Australian region occurred in Torres Strait, northern Australia, in 1995 (31). Three cases (two of which were fatal) were reported from Badu Island in central Torres Strait, 2,000 km from the nearest focus of JE virus activity in Bali. Seroepidemiologic studies showed that the virus was relatively widespread in the central and northern islands with subclinical human cases on four islands and seropositive pigs on nine islands. Ten virus isolates were obtained during the outbreak: two from subclinical human infections (31) and eight from Culex annulirostris mosquitoes collected on Badu Island (32). Sequencing studies showed that these isolates (most closely related to a 1970 isolate from Kuala Lumpur and a 1981 isolate from Bali) (33) were almost identical, suggesting that the outbreak originated from a single source. These studies also showed that all isolates had the same 11 nucleotide deletion in the 3' untranslated region immediately downstream from the stop codon of the open reading frame (34), which provided a signature for comparing any future isolates. After the Badu Island outbreak, inactivated vaccine was offered to all the inhabitants of the northern and central Torres Strait islands (35). During 1996 and 1997, JE virus activity was detected through seroconversions in sentinel pigs on Saibai Island, which is in the north and only about 4 km from the Papua New Guinea coast (36) (J. Lee, D. Phillips, J. Hanna, unpub. results). Seroepidemiologic studies found that virus activity has been widespread in Western Province, Papua New Guinea, since at least 1989, with seropositivity rates of 21% at that time among the Daru-speaking people. Results also showed that seropositivity rates were increasing in the Upper Fly River area and that the virus was spreading geographically (37). Indeed, recent results indicate that JE may have spread to Vanimo on the northern coast by April 1998 and to parts of Milne Bay Province in eastern Papua New Guinea (J. Lee, J. Wangi, P. Siba, G. Tau, unpub. results). The first four clinical cases of JE in Papua New Guinea were observed in 1997 and 1998, with two deaths. All cases were from the Kiunga area of Western Province (J. Oakley, S. Flew, C.A. Johansen, D. Phillips, R.A. Hall, J.S. Mackenzie, unpub. results). Anecdotal evidence suggests that the cases may have resulted in part from the large mosquito numbers associated with the severe drought in 1997. The first JE virus strain isolated in Papua New Guinea was obtained from Cx. annulirostris mosquitoes collected at Lake Murray in Western Province in 1997. Sequence studies have shown that this isolate was almost identical to the 1995 Torres Strait isolates, including the acquisition of the 11 nucleotide deletion (C.A. Johansen, R. Paru, S.A. Ritchie, A. Van den Hurk, M. Bockarie, J.S. Mackenzie, unpub. results). A second outbreak of JE occurred in Torres Strait in March 1998, with one human case from Badu Island and sentinel pigs seroconverting on a number of islands. Shortly afterwards, the first human case on mainland Australia was reported in a fisherman who acquired the infection near the mouth of the Mitchell River in southwest Cape York (38). Extensive seroepidemiologic investigations found no further human infections in communities on Cape York, but domestic pigs had seroconverted both near Mitchell River and near Bamega at the northerly tip of Cape York. Two virus isolations were obtained from pig sera collected near Bamega, and one isolate was obtained from a sentinel pig on Mabuiag Island in Torres Strait (J. Hanna, S. Hills, D. Phillips, J. Lee, unpub. results). Mosquitoes were collected at a number of sites on Cape York as well as on Badu Island. No viruses were isolated from the Cape York mosquitoes, but approximately 44 isolates were obtained from Badu Island—43 from Cx. annulirostris mosquitoes and one from Aedes vigilax mosquitoes (S.A. Ritchie, A. Van Den Hurk, C.A. Johansen, D. Phillips, A. Pyke, J.S. Mackenzie, unpub. results). Nucleotide sequencing studies have shown that the mosquito and pig isolates from Mabuiag and Cape York were closely related to each other, as well as to the 1997 Papua New Guinea Lake Murray and the 1995 Badu Island isolates, including all isolates sharing the 11 base "signature" deletion, which indicated a single virus source for the virus activity in Northern Australia and Papua New Guinea. The focus of activity is probably in Papua New Guinea (C.A. Johansen, A. Drew, D.A. Phillips, A. Pyke, J.S. Mackenzie, unpub. results). JE virus activity in northern Australia began in 1998. Sentinel animal sites are being established to investigate whether the virus has become enzootic in the wildlife. Australia has both the mosquito vectors (Cx. annulirostris) and vertebrate hosts (ardeid birds and pigs) for the virus to become established. In addition, large areas of wetland habitats in Cape York would be conducive to virus enzootic cycles and would increase the potential for the virus to move south to more populous areas of Australia (39). Dengue Viruses Despite a 120-year history, dengue does not appear to be endemic in Australia. Several epidemics over the past decade have been initiated from virus introduced by viremic travelers (27,28,40). Imported cases of dengue in travelers are regularly diagnosed throughout Australia, with 30 to 60 cases reported annually, and growing in number. In most parts of Australia where the local mosquitoes are unable to transmit dengue viruses, these cases pose no risk, but in areas of north Queensland where Ae. aegypti is common and travel between Australia and countries in the Asian-Pacific area is frequent, local transmission and epidemic activity are major risks. The potential for local transmission of dengue viruses is confined to an area of Queensland corresponding to the geographic range of Ae. aegypti, extending from the islands of Torres Strait in the north, to Mount Isa and Boulia in the west, possibly to Roma in the south, and to Gladstone on the east coast (41). Despite this relatively broad geographic range, epidemic activity over the past 2 decades has been restricted from Torres Strait south to Cairns, Townsville, and Charters Towers. The major epidemics over the past 5 years have included a large outbreak of dengue type 2 in 1992 to 1993, principally in Townsville and Charters Towers with more than 2,000 cases, and with the first case of dengue hemorrhagic fever this century (42,43); an outbreak of dengue type 2 in 1996 to 1997 on a number of Torres Strait islands and Cairns, with more than 200 serologically confirmed cases (44,45); and an outbreak of dengue type 3 in 1997 to 1998 largely restricted to Cairns with 239 confirmed cases (46; S. Ritchie, S. Hills, pers. comm.) and also a few cases of dengue type 2. This latter outbreak also included a case of dengue hemorrhagic fever and the first case of dengue encephalopathy in Australia (J. Hanna, unpub. obs.). Nucleotide sequencing of dengue 2 isolates from Australia and a comparison with isolates from elsewhere in the Asian-Pacific region indicated that the 1992-93 isolates were most closely related to an Indonesian virus, whereas the 1996-97 isolates were most closely related to viruses originally isolated in Burkina Faso. This latter finding is of interest because a large outbreak of dengue type 2 occurred on a number of Pacific Islands before and during the Australian outbreak, but the South Pacific viruses were quite distinct from the Australian viruses (45). After the 1992-93 outbreak, a Dengue Fever Management Plan was developed to reduce the potential for epidemic activity from imported cases. The plan has been extremely successful, and a number of imported cases have been recognized early and were contained before they could cause an epidemic (47,48). However, importation, either by continual movement of people between Papua New Guinea and Torres Strait or by movement of people for work, education, or recreation between Papua New Guinea and north Queensland, will always be a major route of entry of the virus. Vector importations occur frequently, with a number of reports for both Ae. aegypti and Ae. albopictus (27), including two recent importations of Ae. albopictus into Townsville in 1997 (49) and Cairns in 1998 (S. Ritchie, pers. comm.). Novel Zoonotic Viral Diseases In the past 4 years, three newly described zoonotic viral diseases have been reported from Australia; two of these diseases are caused by the paramyxoviruses Menangle and Hendra (formerly equine morbillivirus), and the third is caused by Australian bat lyssavirus. Menangle Virus An apparently new virus in the family Paramyxoviridae was isolated from stillborn piglets with deformities at a large commercial piggery in New South Wales (51). The farrowing rate in the piggery decreased from an expected 82% to 60%; the number of live piglets declined in 27% of the litters born; the proportion of mummified and stillborn piglets, some with deformities, increased; and occasional abortions occurred. Virus was isolated from lung, brain, and heart tissues of infected piglets, and shown to be morphologically similar to viruses in the family Paramyxoviridae. No disease was seen in postnatal animals of any age, but a high proportion of sera (>90%) from animals of all ages contained high titers of neutralizing antibodies against the virus. Tests performed at the Australian Animal Health Laboratory confirmed that the virus, named Menangle virus, was unrelated to other known paramyxoviruses, including viruses known to infect pigs 51). Serum from two workers—one at the affected piggery and one at an associated piggery that had received weaned pigs from the original piggery—had high titer, convalescent-phase neutralizing antibodies to the new virus. Both workers had an influenzalike illness with rash during the pig outbreak, but extensive serologic testing showed no evidence of any alternative cause; therefore, the illness is believed to have been caused by the new virus (52). A large breeding colony of gray-headed and little red fruit bats roosted within 200 m of the affected piggery. In a preliminary study, 42 of 125 serum samples collected from fruit bats in New South Wales and Queensland had neutralizing antibodies to the new virus. In addition, antibodies were found in sera collected in 1996, before the outbreak, and from a colony of fruit bats 33 km from the piggery (51). Therefore, the fruit bats are believed to be the primary source of virus causing the outbreak. Sera collected from wild and domestic animals near the affected piggery were seronegative. The geographic range, normal host species, and genetic relationship of this new virus to other paramyxoviruses remain unknown. Nevertheless, Menangle appears to cause fatal disease and malformations in prenatal pigs and may be associated with influenzalike illness in humans. Hendra Virus Hendra virus was first recognized in 1994 after an explosive outbreak of severe, fatal respiratory disease affecting race horses and humans. Twenty race horses in the Brisbane suburb of Hendra were infected; 13 died. The trainer and stable hand were also infected, and the trainer died (53-55). A second incident occurred in Mackay, a coastal town approximately 1,000 km north of Brisbane. Two horses and a farmer died, the latter from severe meningoencephalitis (56-58). The death of the horses and the initial infection of the farmer occurred in 1994 and preceded the Brisbane outbreak; the virus is believed to have then entered a latent phase for 1 year before reactivating to cause fatal encephalitis. No connection was found between the Brisbane and Mackay incidents (56). Experimental studies have shown that in horses and cats, after subcutaneous, intranasal, and oral administration, the virus causes fatal pneumonia (59). In guinea pigs, subcutaneous administration is also fatal, but the infection is more generalized. Black fruit bats (Pteropus alecto) infected by subcutaneous, intranasal, or oral routes contract a subclinical infection and generate an antibody response (M. Williamson, unpub. results). Endothelial cell tropism and formation of syncytia in blood vessels are common pathologic findings in both overt and subclinical infections (B.T. Eaton, M. Williamson, unpub. obs.). Extensive seroepidemiologic studies found no evidence of Hendra virus among horses, other farm animals, or more than 40 species of wildlife in Queensland (56,60; P.L. Young, K. Halpin, H. Field, unpub. results). However, working on the hypothesis that if outbreaks at two distant sites were connected, the most likely wildlife source would be either birds or fruit bats, P.L. Young and colleagues subsequently showed that fruit bats (flying foxes), members of Megachiroptera, were the natural hosts on serologic grounds and by virus isolation, with widespread evidence of infection in four species of fruit bat: the black (Pteropus alecto), grey-headed (P. poliocephalus), little red (P. scapulatus), and spectacled (P. conspicillatus) fruit bats (61,62; P.L. Young et al., unpub. results). Indeed the virus was antigenically and genetically indistinguishable from the earlier horse and human isolates. Thus, it is now clearly established that Hendra virus is a fruit bat virus and is widely distributed throughout the range of pteropid bats in Australia, with serologic evidence of infection in an average of 42% of wild-caught bats, the number of seropositive animals varying with species (53% of 229 P. alecto, 47% of 195 P. poliocephalus, 12% of 115 P. scapulatus, and 41% of 99 P. conspicillatus) and age, but not with geographic distribution (H. Field, unpub. results). Serologic evidence of infection of fruit bats has also been reported from Papua New Guinea. Two species of antibody-positive bats (Dobsonia moluccense, P. neohibernicus) were identified from Madang on the north coast of Papua New Guinea (K. Halpin, H. Field, J.S. Mackenzie, M. Bockarie, P.L. Young, P.W. Selleck, unpub. results), and bats of four more species (D. andersoni, P. capistratus, P. hypomelanus, and P. admiralitatum) were identified in Port Moresby and New Britain (H. Field, S. Hamilton, L. Hall, F. Bornacosso, K. Halpin, P.L. Young, unpub. results). Morphologic features (63) and preliminary sequencing data of the M and F genes (64) suggested that Hendra virus was a member of the Paramyxoviridae, although it had unusual surface projections of two distinct lengths, 15 nm and 18 nm (63). The entire genome of the virus has now been sequenced (65;66; L.F. Wang, B.T. Eaton and colleagues, unpub. results) and has revealed a gene order and P gene organization characteristic of members of the Paramyxovirus and Morbillivirus genera (65). Comparison of its deduced amino acid sequences with those of other family members confirm that Hendra virus is a member of the subfamily Paramyxovirinae, more closely related to members of the Paramyxovirus and Morbillivirus genera than the Rubulavirus genus. Overall, homology with other members of the subfamily is lower than that observed within an individual genus (L.F. Wang, B.T. Eaton, unpub. results). Hendra virus has several distinguishing features, including a genome that is 15% larger than that of other members, with each of the six transcription units containing a very long 3' untranslated region (L.F. Wang, B.T. Eaton, unpub. results). The P/V/C gene has a fourth open reading frame located between those of the C and V proteins and potentially encoding a small basic protein similar to those of some members of the Rhabdoviridae and Filoviridae; its long 3' untranslated region is a common feature of the Filoviridae (65). The sequence of the N gene has also recently been described (66), and like the P/V/C gene, has a 3' untranslated region approximately tenfold longer than other members of the Paramyxovirinae. Although the deduced amino acid sequence of the N protein was slightly more homologous to members of the Morbillivirus genus than to those of other Paramyxovirinae genera, the level of identity was much lower than that observed within the Morbillivirus genus. Three other findings differentiate Hendra from most other members of the Paramyxoviridae: the wide host range (59), the cleavage site of the F protein, and the orientation of the cell surface from which virus is released (B.T. Eaton, W. Michalski, and M. Williamson, unpub. results). An accumulating body of evidence—size of genome, comparative sequence analyses, coding capacity for a small basic protein in the P gene, morphologic features, host range, and various biologic properties, together with the wildlife host of the virus—suggests that the virus had been misnamed—it was neither an equine virus nor a morbillivirus—although the name was relevant when the virus was first isolated. It has therefore been suggested that the virus be renamed Hendra and be classified in a new genus within the Paramyxovirinae (59,65,66). A number of aspects of the ecology of the virus remain to be determined. For instance, despite the obvious ubiquity of the virus in the fruit bat population and the extremely close relationship between bat caregivers and bats, there is no evidence of seroconversions among caregivers, despite their close contact with up to 1,000 bats per year (50). Specimens of persons who have died of either pneumonia or encephalitis of unknown etiology were virus-negative (C. Allan, J.S. Mackenzie, L.A. Selvey, unpub. results). Furthermore, all human infections appear to have been transmitted by horses. Thus, the virus appears to have low transmissibility to humans; it also appears to be linked with pregnancy: the index case of the Brisbane outbreak was a pregnant mare, a pregnant mare was involved in the Mackay incident, both incidents occurred during the birthing season of flying foxes, and virus was first recovered from uterine fluid of a pregnant animal (3). Thus, a number of questions remain about the ecologic, biologic, and pathologic characteristics of Hendra virus: 1) the infectivity and virulence of the virus and why it seems extremely difficult to transmit naturally between and within some susceptible host species, 2) classification of the virus, 3) role of pregnancy to transmission of the virus, 4) role of prior infection in horses in human infection, 5) method of transmission between fruit bats and from fruit bats to horses, 6) tropism of the virus, and 7) potential for producing a latent infection in humans. Australian Bat Lyssavirus Australia had been considered free of rabies and rabieslike viruses until 1996 when a new lyssavirus, closely related to classic rabies virus, was first identified in a fixed brain specimen from a young black flying fox (P. alecto), with unusual neurologic symptoms. Since this original isolation, a further 42 isolates have been obtained from all four species of fruit bat, with most isolates from black and little red (P. scapulatus) flying foxes, and four isolates from an insectivorous bat (Microchiroptera), the yellow-bellied sheathtail bat (Saccolaimus flavicentris) (P. Daniels, R. Lunt, unpub. results). The isolates were from as far apart as Melbourne and Darwin, but most were from Queensland. Antibodies to rabies virus (REFIT assay) have been detected in 2.6% of 345 bat nonrandom samples submitted to the Australian Animal Health Laboratory (P. Daniels, R. Lunt, unpub. results). Antibody has been detected in both infected and apparently healthy bats, but whether this reflects the ability of bats to recover from infection or become latently infected with the virus is not known. Analysis using nucleocapsid-specific monoclonal antibodies showed a strong relationship between this new lyssavirus and serotype 1 rabies virus (67). Indeed, rabies vaccine may elicit a protective immune response in humans, indicating the antigenic similarity of Australian bat lyssavirus and classic rabies virus (P.K. Murray, pers. comm. to the Lyssavirus Expert Committee [68]). Phylogenetic studies of the N protein sequences indicated that the Australian virus was genetically distinct from classic rabies (genotype 1) and was, therefore, a previously unrecognized member of the Lyssavirus genus and represented a new genotype, genotype 7 (67). Two human infections have been attributed to Australian bat lyssavirus. One fatal rabieslike infection was in a female bat caregiver from Rockhampton, Queensland (69,70). An isolate of Australian bat lyssavirus obtained post-mortem was antigenically and genetically similar to the virus from the insectivorous yellow-bellied sheathtail bat (A.R. Gould, R. Lunt, P. Daniels, unpub. results). A second death has recently been reported in Queensland (J. Hann, J. Faoagali, G. Smith, I Serafin, J. Northill, unpub. obs.). The infection was in a 27-year-old woman from Mackay, who died 2 years after a bat bite (by a large flying fox). Polymerase chain reaction (PCR) testing of RNA extracted from saliva and nuchal biopsy proved vital to the antemortem diagnosis. Immunofluorescence staining of postmortem samples confirmed the diagnosis. Preliminary sequencing of the amplicon has indicated that the virus is very similar to other lyssaviruses isolated from flying foxes but clearly distinct from a virus isolate from a yellow-bellied sheathtail bat (I. Serafin, G. Smith, J. Hanna, B. Harrower, J. Northill, A. Westcott, unpub. obs.). More extensive sequencing of the human and bat isolates is under way. Measures to prevent further human infection have been implemented (68,71). As with Hendra virus, a number of questions remain about the ecology and biology of Australian bat lyssavirus. The finding of well, antibody-positive bats, which suggests that bats can either recover from infection or that they can be silently infected, needs to be investigated, particularly with respect to infectivity and possible transmissibility. More information is needed on the geographic and host range of the virus ecology within bat communities and risk for transmission to terrestrial animals. These novel zoonotic viruses appear to have frugivorous bats as their natural vertebrate hosts. While little is known of the viral fauna of fruit bats (or indeed of most wildlife species) in Australia, the occurrence of these three zoonotic viruses from bats over 3 years suggests that further prospective studies of diseases of wildlife are warranted. Indeed, two paramyxoviruslike viruses unrelated to any other known paramyxoviruses (K. Halpin, P.L. Young, unpub. results) have recently been isolated from flying foxes. Conclusions The vector-borne and zoonotic diseases in this editorial encompass three patterns of emergence: known diseases increasing in incidence or geographic range (e.g., dengue and JE virus, respectively); new infectious agents as etiologic agents of known diseases (e.g., Australian bat lyssavirus as a cause of a rabieslike illness); and new infectious agents causing previously unrecognized diseases (e.g., Hendra virus). All three patterns demonstrate the need (and international responsibility) for ongoing surveillance and monitoring. In Australia, surveillance is the legislative responsibility of the individual states and territories. A Communicable Diseases Network Australia-New Zealand was established in 1989 to improve the control of communicable diseases in Australia by coordinating national surveillance activities and responses to outbreaks and by training public health staff. In 1996, Australia developed a National Communicable Diseases Surveillance Strategy to provide a national framework to monitor infectious diseases and plan and prioritize interventions. Components of the strategy include improvements to the national surveillance infrastructure, better monitoring of diseases and surveillance data, and better response to outbreaks. The strategy is being implemented and may provide the mechanism for a national response to new and reemerging diseases. Acknowledgments I thank my many colleagues—Bryan Eaton, Lin-Fa Wang, Peter Daniels, Ross Lunt, Jeffrey Hanna, Scott Ritchie, Peter Young, Kim Halpin, Greg Smith, Debbie Phillips, Cheryl Johansen, Hume Field, Steve Flew, and John Oakley—whose unpublished work I have been permitted to quote. Address for correspondence: John S. Mackenzie, Department of Microbiology, The University of Queensland, Brisbane, Qld 4072, Australia; fax: 61-7-3365-4620; e-mail: jmac@biosci.uq.edu.au. References 1. Beaman MH. Emerging infections in Australia. Ann Acad Med Singapore 1997;26:609-15. 2. Longbottom H. Emerging infectious diseases. Commun Dis Intell 1997;21:89-93. 3. Mackenzie JS, Bolton W, Cunningham AL, Frazer IH, Gowans EJ, Grohmann GS, et al. Emerging viral diseases of humans: an Australian and New Zealand perspective. In: Asche V, editor. Recent advances in microbiology. Vol. 5. Melbourne: Australian Society for Microbiology Inc.; 1997. p. 13-130. 4. Mackenzie JS. Emerging viral diseases: some comments from a regional perspective. P N G Med J. In press 1998. 5. Della-Porta AJ. Emerging and re-emerging diseases of animals in Australia. In: Asche V, editor. Recent advances in microbiology. Vol. 5. Melbourne: Australian Society for Microbiology Inc.; 1977. p. 131-201. 6. Desmarchelier PM. Foodborne disease: emerging problems and solutions. Med J Aust 1996;165:668-71. 7. Crerar SK, Dalton CB, Longbottom HM, Kraa E. Foodborne disease: current trends and future surveillance needs in Australia. Med J Aust 1996;165:672-5. 8. Collignon PJ. Antibiotic resistance: is it leading to the re-emergence of many infections of the past? In: Asche V, editor. Recent advances in microbiology. Vol. 5. Melbourne: Australian Society for Microbiology Inc.; 1997. p. 203-56. 9. Collignon PJ, Bell JM. Drug-resistant Streptococcus pneumoniae: the beginning of the end for many antibiotics? Australian Group on Antimicrobial Resistance. Med J Aust 1996;164:64-7. 10. Heath CH, Blackmore TK, Gordon DL. Emerging resis-tance in Enterococcus spp. Med J Aust 1996;164:116-20. 11. Maguire GP, Arthur AD, Boustead PJ, Dwyer B, Currie BJ. Emerging epidemic of community-acquired methicillin-resistant Staphylococcus aureus infection in the Northern Territory. Med J Aust 1996;164:721-3. 12. Gratten M, Nimmo G, Carlisle J, Schooneveldt J, Seneviratne E, Kelly R, et al. Emergence of further sero-types of multiple drug-resistant Streptococcus pneumoniae in Queensland. Commun Dis Intell 1997;21:133-6. 13. Dawson D. Tuberculosis in Australia: bacteriologically confirmed cases and drug resistance, 1996. Report of the Australian Mycobacterium Reference Laboratory Network. Commun Dis Intell 1998;22:183-7. 14. Givney R, Vickery A, Holliday A, Pegler M, Benn R. Evolution of an endemic methicillin-resistant Staphylococcus aureus population in an Australian hospital from 1967 to 1996. J Clin Microbiol 1998;36:552-6. 15. Maguire GP, Arthur AD, Boustead PJ, Dwyer B, Currie BJ. Clinical experience and outcomes of community-acquired and nosocomial methicillin-resistant Staphylococcus aureus in a northern Australian hospital. J Hosp Infect 1998;38:273-81. 16. Grimwood K, Collignon PJ, Currie BJ, Ferson MJ, Gilbert GL, Hogg GG, et al. Antibiotic management of pneumococcal infections in an era of increased resistance. J Paediatr Child Health 1997;33:287-95. 17. Patel MS, Collignon PJ, Watson CR, Condon RJ, Doherty RR, Merianos A, et al. New guidelines for management and prevention of meningococcal disease in Australia. Meningococcal Disease Working Party of the National Health and Medical Research Council. Med J Aust 1997;166:598-601. 18. Cameron S, Walker C, Beers M, Rose N, Anear E. Enterohaemorrhagic Escherichia coli outbreak in South Australia associated with the consumption of mettwurst. Commun Dis Intell 1995;19:70-1. 19. Ng S, Rouch G, Dedman R, Harries B, Boyden A, McLennan L, et al. Human salmonellosis in peanut butter. Commun Dis Intell 1996;20:326-7. 20. Hook D, Jalaludin B, Fitzsimmons G. Clostridium perfringens food-borne outbreak: an epidemiological investigation. Aust N Z J Public Health 1996;20:119-22. 21. Stafford R, Strain D, Heymer M, Smith C, Trent M, Beard J. An outbreak of Norwalk virus gastroenteritis following consumption of oysters. Commun Dis Intell 1997;21:317-20. 22. Salmonellosis outbreak. Commun Infect Dis 1998;22:155. 23. Lemmon JM, McAnulty JM, Bawden-Smith J. Outbreak of cryptosporidiosis linked to an indoor swimming pool. Med J Aust 1996;165:613-6. 24. Cryptosporidiosis outbreak. Commun Dis Intell 1998;22:22. 25. Parasites in water. Commun Dis Intell 1998;22:190. 26. Hyland CA, Mison L, Solomon N, Cockerill J, Wang L, Hunt J, et al. Exposure to GBV-C/HGV in selected Australian adult and children populations. Transfusion. In press 1998. 27. Mackenzie JS, Broom AK, Hall RA, Johansen CA, Lindsay MD, Phillips DA, et al. Arboviruses in the Australian region, 1990 to 1998. Commun Dis Intell 1998;22:93-100. 28. Mackenzie JS, Lindsay MD, Coelen RJ, Broom AK, Hall RA, Smith DW. Arboviruses causing human disease in the Australasian zoogeographic region. Arch Virol 1994;136:447-67. 29. Lindsay MDA, Johansen CA, Broom AK, Smith DW, Mackenzie JS. Emergence of Barmah Forest virus in Western Australia. Emerg Infect Dis 1995;1:22-6. 30. Lindsay MD, Johansen CA, Smith DW, Wallace MJ, Mackenzie JS. An outbreak of Barmah Forest virus disease in the south-west of Western Australia. Med J Aust 1995;162:291-4. 31. Hanna J, Ritchie S, Phillips DA, Shield J, Bailey MC, Mackenzie JS, et al. An outbreak of Japanese encephalitis in the Torres Strait, Australia, 1995. Med J Aust 1996;256-60. 32. Ritchie SA, Phillips D, Broom A, Mackenzie J, Poidinger P, Van Den Hurk A. Isolation of Japanese encephalitis virus from Culex annulirostris mosquitoes in Australia. Am J Trop Med Hyg 1997;56:80-4. 33. Mackenzie JS, Poidinger M, Phillips D, Johansen C, Hall RA, Hanna J, et al. Emergence of Japanese encephalitis virus in the Australasian region. In: Saluzzo JF, Dodet B, editors. Factors in the emergence of arboviral diseases. Paris: Elsevier; 1997. p. 191-201. 34. Poidinger M, Hall RA, Mackenzie JS. Molecular characterisation of the Japanese encephalitis serocomplex of the Flavivirus genus. Virology 1996;218:417-21. 35. Hanna J, Barnett D, Ewaid D. Vaccination against Japanese encephalitis in the Torres Strait. Commun Dis Intell 1996;20:188-90. 36. Shield J, Hanna J, Phillips D. Reappearance of the Japanese encephalitis virus in the Torres Strait, 1996. Commun Dis Intell 1996;20:191. 37. Johansen C, Ritchie S, Van Den Hurk A, Bockarie M, Hanna J, Phillips D, et al. The search for Japanese encephalitis virus in the Western province of Papua New Guinea. In: Kay BH, Brown MD, Aaskov JG, editors. Arbovirus research in Australia. Vol. 7. Brisbane: Queensland Institute of Medical Research; 1997. p. 131-6. 38. Japanese encephalitis on the Australian mainland. Commun Dis Intell 1998;22:60. 39. Mackenzie JS. Japanese encephalitis: an emerging disease in the Australian region, and its potential risk to Australia. In: Kay BH, Brown MD, Aaskov JG, editors. Arbovirus research in Australia. Vol. 7. Brisbane: Queensland Institute of Medical Research; 1997. p. 166-70. 40. Mackenzie JS, LaBrooy JT, Hueston L, Cunningham AL. Dengue in Australia [editorial]. J Med Microbiol 1996;45:159-61. 41. Sinclair DP. The distribution of Aedes aegypti in Queensland, 1990 to 30 June 1992. Commun Dis Intell 1992;16:400-3. 42. Phillips D, Pearce M, Weimers M, Blumke G. Dengue 2 infection in northern Queensland. Commun Dis Intell 1992;16:192-3. 43. Row D, Pearce M, Hapgood G, Sheridan J. Dengue and dengue haemorrhagic fever in Charters Towers, Queensland. Commun Dis Intell 1993;17:182-3. 44. Griffiths M, Ritchie S, Terry D, Norton R, Phillips D. An outbreak of dengue 2 in the Torres Strait. Commun Dis Intell 1997;21:33. 45. Hanna JN, Ritchie SA, Merritt AD, Van den Hurk AF, Phillips DA, Serafin IL, et al. Two contiguous outbreaks of dengue type 2 in north Queensland. Med J Aust 1998;168:221-5. 46. Dengue 3 in Cairns: the story so far. Commun Dis Intell 1998;22:109-10. 47. Ritchie S, Hanna J, Van den Hurk A, Harley D, Lawrence R, Phillips D. Importation and subsequent local transmission of dengue 2 in Cairns. Commun Dis Intell 1995;19:366-70. 48. Hanna J, Ritchie S, Tiley S, Phillips D. Dengue imported from Papua New Guinea. Commun Dis Intell 1995;19:447. 49. Foley P, Hemslaey C, Muller K, Maroske G, Ritchie S. Importation of Aedes albopictus in Townsville, Queensland. Commun Dis Intell 1998;22:3-4. 50. Selvey L, Taylor R, Arklay A, Gerrard J. Screening of bat carers for antibodies to equine morbillivirus. Commun Dis Intell 1996;20:477-8. 51. Philbey AW, Kirkland PD, Ross AD, Davies RJ, Gleeson AB, Love RJ, et al. An apparently new virus (family Paramyxoviridae) infectious for pigs, humans, and fruit bats. Emerg Infect Dis 1998;4:269-71. 52. Chant K, Chan R, Smith M, Dwyer DE, Kirkland P, the NSW Expert Group. Probable human infection with a newly described virus in the family Paramyxoviridae. Emerg Infect Dis 1998;4:273-5. 53. Murray K, Rogers R, Selvey L, Selleck P, Hyatt A, Gould A, et al. A novel morbillivirus pneumonia of horses and its transmission to humans. Emerg Infect Dis 1995;1:31-3. 54. Murray K, Selleck P, Hooper P, Hyatt A, Gould A, Gleeson L, et al. A morbillivirus that caused fatal disease of horses and humans. Science 1995;268:94-6. 55. Selvey LA, Wells RM, McCormack JG, Ansford AJ, Murray K, Rogers RJ, et al. Infection of humans and horses by a newly described morbillivirus. Med J Aust 1995;162:642-5. 56. Rogers RL, Douglas IC, Baldock FC, Glanville RJ, Seppanen KT, Gleeson LJ, et al. Investigation of a second focus of equine morbillivirus infection in coastal Queensland. Aust Vet J 1996;74:243-4. 57. O'Sullivan JD, Allworth AM, Paterson DL, Snow TM, Boots R, Gleeson LJ, et al. Fatal encephalitis due to a novel paramyxovirus transmitted from horses. Lancet 1997;349:93-5. 58. Hooper PT, Gould AR, Russell GM, Kattenbelt JA, Mitchell G. The retrospective diagnosis of a second outbreak of equine morbillivirus infection. Aust Vet J 1996;74:244-5. 59. Murray K, Eaton B, Hooper P, Wang L, Williamson M, Young P. Flying foxes, horses, and humans: a zoonosis caused by a new member of the Paramyxoviridae. In: Scheld WM, Armstrong D, Hughes JM, editors. Emerging infections 1. Washington: American Society for Microbiology Press; 1998. p. 43-58. 60. Ward MP, Black PF, Childs AJ, Baldock FC, Webster WR, Rodwell BJ, et al. Negative findings from serological studies of equine morbillivirus in the Queensland horse population. Aust Vet J 1996;74:241-3. 61. Young PL, Halpin K, Selleck PW, Field H, Gravel JL, Kelly MA, et al. Serologic evidence for the presence in pteropus bats of a paramyxovirus related to equine morbillivirus. Emerg Infect Dis 1996;2:239-40. 62. Young P, Halpin K, Field H, Mackenzie J. Finding the wildlife reservoir of equine morbillivirus. In: Asche V, editor. Recent advances in microbiology. Vol. 5. Melbourne: Australian Society for Microbiology Inc.; 1997. p. 1-12. 63. Hyatt AD, Selleck PW. Ultrastructure of equine morbillivirus. Virus Res 1996;43:1-15. 64. Gould AR. Comparison of the deduced matrix and fusion protein sequences of equine morbillivirus with cognate genes of the Paramyxoviridae. Virus Res 1996;43:17-31. 65. Wang LF, Michalski WP, Yu M, Pritchard LI, Crameri G, Shiell B, et al. A novel P/V/C gene in a new member of the Paramyxoviridae family, which causes lethal infection in humans, horses, and other animals. J Virol 1998;72:1482-90. 66. Yu M, Hansson E, Shiell B, Michalski W, Eaton BT, Wang LF. Sequence analysis of the Hendra virus nucleoprotein gene: comparison with other members of the subfamily Paramyxoviridae. J Gen Virol 1998;79:1775-80. 67. Gould AR, Hyatt AD, Lunt R, Kattenbelt JA, Hengstberger S, Blacksell SD. Characterisation of a novel lyssavirus isolated from Pteropid bats in Australia. Virus Res 1998;54:165-87. 68. Lyssavirus Expert Group. Prevention of human lyssavirus infection. Commun Dis Intell 1996;20:505-7. 69. Hooper PT, Lunt RA, Gould AR, Samaratunga H, Hyatt AD, Gleeson LJ, et al. A new lyssavirusthe first endemic rabies-related virus recognized in Australia. Bulletin de l'Institut Pasteur 1997;95:209-18. 70. Allworth A, Murray K, Morgan J. A case of encephalitis due to a lyssavirus recently identified in fruit bats. Commun Dis Intell 1996;20:504. 71. Lyssavirus Expert Group. Update on bat lyssavirus. Commun Dis Intell 1996;20:535. ——————————————————————————————————————————————————————————————————————————— Perspectives ——————————————————————————————————————————————————————————————————————————— Perspectives The Economic Impact of Staphylococcus aureus Infection in New York City Hospitals Robert J. Rubin, Catherine A. Harrington, Anna Poon, Kimberly Dietrich, Jeremy A. Greene, and Adil Moiduddin The Lewin Group, Fairfax, Virginia, USA --------------------------------------------------------------------------- We modeled estimates of the incidence, deaths, and direct medical costs of Staphylococcus aureus infections in hospitalized patients in the New York City metropolitan area in 1995 by using hospital discharge data collected by the New York State Department of Health and standard sources for the costs of health care. We also examined the relative impact of methicillin-resistant versus -sensitive strains of S. aureus and of community-acquired versus nosocomial infections. S. aureus-associated hospitalizations resulted in approximately twice the length of stay, deaths, and medical costs of typical hospitalizations; methicillin-resistant and -sensitive infections had similar direct medical costs, but resistant infections caused more deaths (21% versus 8%). Community-acquired and nosocomial infections had similar death rates, but community-acquired infections appeared to have increased direct medical costs per patient ($35,300 versus $28,800). The results of our study indicate that reducing the incidence of methicillin-resistant and -sensitive nosocomial infections would reduce the societal costs of S. aureus infection. Each year approximately two million hospitalizations result in nosocomial infections (1). In a study of critically ill patients in a large teaching hospital, illness attributable to nosocomial bacteremia increased intensive care unit stay by 8 days, hospital stay by 14 days, and the death rate by 35% (2). An earlier study found that postoperative wound infections increased hospital stay an average of 7.4 days (3). Staphylococcus aureus was the most common cause of nosocomial infections reported in the National Nosocomial Surveillance System between 1990 to 1996 (4). The leading cause of nosocomial pneumonia and surgical site infections and the second leading cause of nosocomial bloodstream infections (4), S. aureus also causes community-acquired infections (e.g., osteomyelitis and septic arthritis, skin infections, endocarditis, and meningitis). More than 95% of patients with S. aureus infections worldwide do not respond to first-line antibiotics such as penicillin or ampicillin (5). Additionally, methicillin-resistant strains of S. aureus (MRSA) are common. First reported in the 1960s (6), MRSA has become increasingly prevalent since the 1980s (7,8) and is now endemic in many hospitals and even epidemic in some, with resistance in approximately 30% of all S. aureus infections (8). Vancomycin is the only drug that can consistently treat MRSA. However, beginning in 1989, hospitals have reported a rapid increase in vancomycin resistance in enterococci (VRE) (9). Increased vancomycin use helps select for VRE, and even a small increase in incidence of VRE infection could lead to cross-resistance in S. aureus, since genes conferring vancomycin resistance might be transferred from VRE (10). In 1996, Japan reported the first case of S. aureus infection with intermediate resistance to vancomycin (11). In 1997, two unrelated cases of S. aureus infection with intermediate resistance to vancomycin were reported in the United States (Michigan and New Jersey) (12). In both cases, patients had been treated with multiple courses of vancomycin for repeated MRSA infections over the 6 months before the S. aureus infection with intermediate resistance to vancomycin; additionally, VRE colonization had been diagnosed 7 months before the S. aureus infection with intermediate resistance to vancomycin in the New Jersey patient. The emergence of S. aureus infection with intermediate resistance to vancomycin in the United States suggests that S. aureus strains are constantly evolving and full resistance may develop (12). The various ways of controlling MRSA (13) are still being debated. The elimination of endemic MRSA in hospitals is difficult and costly (14-17). In general, infection control in the United States is less stringent than in Canada and in some European countries, where identification of known carriers, prospective surveillance of patients and hospital workers, and use of nasal mupirocin have helped control drug-resistant S. aureus infection rates (18). Knowledge of the scope of the problem is helpful for hospital administrators, insurers, and medical personnel who make policy decisions on control measures to prevent the spread of MRSA and the emergence of vancomycin-resistant S. aureus. However, the economic cost of S. aureus infections is not well known. Many studies focus on the cost of nonorganism-specific nosocomial infections (2,19,20). Moreover, the reported cost of a nosocomial infection varies because of the wide range of study populations, sites of infection, and methods used (16,21). The few investigations into the cost of S. aureus infections have focused on the differential cost of MRSA and MSSA infections (22,23) and are case studies of outbreaks in single hospitals. Thus, they do not provide perspective on the scope of the problem for a population over time. We estimated the incidence, death rate, and cost of S. aureus infections associated with hospitalization in the New York City metropolitan area in 1995. We selected this geographic region because of its high prevalence of multidrug-resistant infections (24,25). We also compared the relative contributions of nosocomial versus community-acquired infections and methicillin-sensitive (MSSA) versus methicillin-resistant S. aureus. The Study Data The 1995 Statewide Planning and Research Cooperative System (SPARCS) Administratively Releasable File was the primary source of data (26). SPARCS is a database of all hospital discharges in New York state, as reported by hospitals to the State of New York Department of Health, and the Administratively Releasable File contains discharge information on hospital location, patient characteristics (age, sex, race, ethnicity), and visit characteristics (primary diagnosis, secondary diagnoses, primary procedure, secondary procedures, length of stay, total charges, patient status, and disposition). We analyzed data for hospitals in the following New York City metropolitan area counties: Bronx, Dutchess, Kings, Manhattan, Nassau, Orange, Putnam, Queens, Richmond, Rockland, Suffolk, Ulster, and Westchester. Data on infection incidence or resource use not in SPARCS were obtained through a comprehensive literature search or estimated by a clinical panel consisting of four physicians specializing in infectious disease. Other sources for cost information were the 1995 Medicare Fee Schedule (27) for physician fees and the 1995 Red Book (28) for outpatient pharmaceutical average wholesale prices. Definitions We identified patients with the most common types of hospital-associated S. aureus infections: pneumonia, bacteremia, endocarditis, surgical site infections, osteomyelitis, and septic arthritis (Table 1) from SPARCS, which uses the International Classification of Diseases, Ninth Revision, Clinical Modification (ICD-9-CM) diagnosis codes (29). With the exception of ICD-9-CM code 482.4 (staphylococcal pneumonia) and 038.1 (staphylococcal septicemia), these codes are not organism-specific. Table 1. ICD-9-CM codes used to identify infections in Statewide Planning and Research Cooperative System ------------------------------------------------------------------------------ Type of ICD-9-CM(sup a) Infection Description ------------------------------------------------------------------------------ Pneumonia 482.4 Pneumonia due to staphylococcus Bacteremia 038.1 Staphylococcal septicemia 790.7 Bacteremia 996.62 Infection and inflammatory reaction due to internal vascular device, implant, and graft Endocarditis 421.0 Acute and subacute bacterial endocarditis 996.61 Infection and inflammatory reaction due to cardiac device, implant, and graft Surgical site 998.3 infection Disruption of operation wound 998.5 Postoperative infection Osteomyelitis 730.01-730.09 Acute osteomyelitis 730.10-730.19 Chronic osteomyelitis Septic arthritis 711.00-711.09 Pyogenic arthritis 996.66 Infection and inflammatory reaction due to internal joint problems ------------------------------------------------------------------------------ (sup a)International classification of diseases, 9th Revision, Clinical Modification, 1995. To identify S. aureus infections, we used the nonorganism–specific codes in conjunction with an additional ICD-9-CM code to identify the bacterial agent (i.e., 041.11 bacterial infection due to S. aureus in conditions classified elsewhere and of unspecified site). Patients with multiple infections were counted only once in the overall incidence rate. Their primary or first occurrence of a diagnosis of interest was used. Because source of infection (nosocomial versus community-acquired) is not reported in SPARCS, we assumed that specific types of disease were either nosocomial or community-acquired on the basis of the clinical panel opinion (Table 2). Table 2. Definitions of nosocomial or community-acquired Staphylococcus aureus infections ------------------------------------------------------------------------------ Type of Infection Nosocomial Community-acquired ------------------------------------------------------------------------------ Pneumonia Secondary diagnosis(sup a) Primary diagnosis(sup a) Bacteremia Catheter- or surgery- Noncatheter- and nonsurgical- associated infections(sup b)associated infections Endocarditis Prosthetic valve infections Natural valve infections Surgical site infection (SSI) All SSIs None Osteomyelitis None All Septic arthritis Prosthetic joint infections Natural joint infections ------------------------------------------------------------------------------ (sup a)ICD-9-CM 482.4 as the primary diagnosis vs. 482.4 as one of several other diagnoses. (sup b)ICD-9-CM 996.62, or 038.1 associated with a surgical ICD procedure code, or 790.7 associated with a surgical ICD procedure code. Modeling the Incidence Rate ICD-9-CM code 041.11 (bacterial infection due to S. aureus) is not widely used by reporting hospitals. Therefore, the incidence of S. aureus infections based on the counts of 041.11 in SPARCS would underestimate the number of cases. We estimated the incidence of S. aureus infections (except pneumonia) as follows (Table 3): the total incidence of each type of infection (e.g., endocarditis) in SPARCS was multiplied by the estimated percentage attributable to S. aureus (determined by research or clinical panel opinion) to give the total number of infections due to S. aureus. The incidence of pneumonia was equated with the occurrence of the ICD-9-CM code 482.4 (staphylococcal pneumonia). For ICD-9-CM code 038.1 (staphylococcal septicemia), we assumed that only 50% of infections were attributable to S. aureus (with the remainder attributable to S. epidermidis) (30). Table 3. Incidence of Staphylococcus aureus infections from research or clinical panel ------------------------------------------------------------------------------- Type of infection Description S. aureus % Reference ------------------------------------------------------------------------------- Bacteremia Staphylococcal septicemia 50 30 Bacteremia 15 31,32 Infection and inflammatory 16 4 reaction due to internal vascular device, implant, and graft Endocarditis Acute and subacute bacterial Clinical endocarditis 30 panel Infection and inflammatory 14 33 reaction due to cardiac device, implant, and graft Surgical site Disruption of operation wound 20 4 infection and postoperative infection Osteomyelitis Acute and chronic osteomyelitis 50 34,35 Septic 33 arthritis Pyogenic arthritis 11 (age <5 yr) 33 (age 5-18 yr) 55 (age >18 yr) Infection and inflammatory 25 33 reaction due to internal joint prosthesis ------------------------------------------------------------------------------- Modeling Death Rates The death rates attributable to bacteremia, endocarditis, or community-acquired pneumonia were assumed to be equal to the death rates found when these infections were coded as a primary diagnosis in SPARCS and 041.11 was used as a secondary diagnosis. For nosocomial pneumonia, however, we assumed that the attributable death rate was a percentage of the actual death rate——for ventilator-associated pneumonia patients, death rate is a function of both the severity of underlying disease and the pneumonia. A series of matched-cohort studies have demonstrated that the death rate attributable to ventilator-associated pneumonia is 0% to 57% of the actual death rate (36-39). On the basis of this research and expert panel judgment, nosocomial pneumonia in ventilator-associated pneumonia patients (identified by ICD-9-CM V46.0 or V46.1) was assumed to have an attributable death rate of 50% of the death rate found in SPARCS (30,40). We assumed that the attributable death rate of nonventilator-associated pneumonia was the death rate found in SPARCS. On the basis of the low death rate found in SPARCS (approximately 2%), we assumed that no deaths were attributable to osteomyelitis, septic arthritis, or surgical site infections. Modeling Direct Medical Costs Direct medical costs were defined as hospital costs attributable to S.aureus infection, professional fees incurred during hospitalization, and costs of other infection-related medical services provided after discharge. For each infection, total direct medical costs were calculated by multiplying the average direct medical cost per patient by the incidence of disease. Average hospital costs attributable to S. aureus per patient were assumed to be equal to the average hospital charge from SPARCS when the infection (e.g., pneumonia, bacteremia) was coded as a primary diagnosis and 041.11 was used as a secondary diagnosis. Professional fees incurred during hospitalization include physician visits and consultations for evaluation and management, as well as radiologic, surgical, and anesthesiologic costs. The average frequency of physician services per patient was based on clinical panel estimates. Costs of these services were based on 1995 Medicare Payment Rates for the Long Island, New York, area as an intermediate point between New York City costs and those of outlying counties. Costs of medical services after discharge include those of postdischarge complications (e.g., abscesses, aneurysms) requiring rehospitalization, home-based intravenous antibiotic therapy, and outpatient oral antibiotic therapy. The average frequency of other medical services provided per patient was based on clinical panel estimates. Costs of hospital readmission were based on SPARCS charges; costs of home-based intravenous therapy were based on literature estimates (40,41); and costs of outpatient medications were based on average wholesale prices (25). Modeling MRSA and MSSA S. aureus Infections SPARCS does not identify MRSA or MSSA infections, and a code for infection with a drug-resistant organism (V09) is rarely used. Therefore, we modeled the comparative incidence, death rate, and cost of MRSA and MSSA. We computed the incidence of MRSA and MSSA infections by using the estimate that 29% of infections were due to MRSA (8). The clinical panel estimated that 10% of community-acquired infections were due to MRSA (includes infections acquired at long-term care facilities). The number of deaths for MRSA and MSSA infections was estimated as follows: the clinical panel estimated a risk ratio for death rates of MRSA and MSSA infections, and deaths due to MRSA and MSSA infections were calculated from the estimated risk ratio and the overall number of deaths due to S. aureus infection. We estimated the direct medical cost per patient for MRSA and MSSA infections as follows: differences in resource use for those with MRSA and MSSA infections were identified by the clinical panel; these differences were converted into differences in cost using a method similar to that described above for modeling direct medical costs; and average costs for MRSA and MSSA infections were calculated by using the average cost for an S. aureus infection and the average difference in cost between MRSA and MSSA infections. Incidence, Death Rate, and Attributable Costs S. aureus Infection Of 1,351,362 nonobstetrical hospital discharges in SPARCS for New York City in 1995, an estimated 13,550 (1.0%) were discharges of patients with S. aureus infections (Table 4). The total direct medical costs incurred by these patients was an estimated $435.5 million—average length of stay nearly 20 days, direct cost of infection, $32,100 (Table 4). The number of deaths was estimated at 1,400 (a 10% death rate). In contrast, the hospital charges for the average hospital stay in SPARCS (for all nonobstetrical discharges) were $13,263—average length of stay 9 days, death rate 4.1%. Thus, patients with S. aureus infection had approximately twice the cost, length of stay, and death rate of a typical hospitalized patient. Table 4. Incidence, length of stay, costs, and death rates of Staphylococcus aureus infections by type of infection ------------------------------------------------------------------------------- Direct Medical Costs ----------------------- Length of Deaths stay Total Per patient Type of infection Incidence (days) ($M) ($) Total % ------------------------------------------------------------------------------- Pneumonia 3,600 22.2 128.3 35,400 890 25 Bacteremia 4,400 18.0 137.0 31,300 470 11 Endocarditis 550 25.9 25.8 47,200 40 7 Surgical site ND(sup a) infection 2,300 13.6 50.5 21,800 ND Osteomyelitis 2,000 23.9 68.4 35,000 ND ND Septic arthritis 700 22.0 25.5 35,100 ND ND Total or average 13,550 19.8 435.5 32,100 1,400 10 ------------------------------------------------------------------------------- (sup a)ND=no data. Pneumonia and bacteremia represented most S. aureus infections and accounted for 60% of the total direct medical costs and 97% of the number of deaths. Endocarditis caused the longest stay (26 days) and highest direct cost per patient ($47,200); surgical site infection caused the shortest stay (14 days) and lowest direct cost per patient ($21,810). Hospital charges were an average of $29,000 (90% of the total costs); professional fees were an average of $2,300 (7%); and postdischarge costs represented $800 (3%) (Table 5). Table 5. Direct medical charges—average hospital facility charges, professional fees, and postdischarge costs per case ---------------------------------------------------------------------- Profes- Post- Hospital sional discharge charges fees costs Type of infection $ (%) $ (%) $ (%) Total $ ---------------------------------------------------------------------- Pneumonia 33,400 (94) 2,000 (6) ND(sup a) 35,400 Bacteremia 27,900 (89) 2,100 (7) 1,300 (4) 31,300 Endocarditis 41,700 (88) 4,300 (9) 1,200 (3) 47,200 Surgical site infection 20,200 (93) 1,600 (7) ND 21,800 Osteomyelitis 30,000 (86) 3,200 (9) 1,800 (5) 35,000 Septic arthritis 30,600 (87) 3,100 (9) 1,400 (4) 35,100 Average 29,000 (90) 2,300 (7) 800 (3) 32,100 ---------------------------------------------------------------------- (sup a)ND=no data. Nosocomial Infection Nosocomial infections accounted for 46% of the total incidence of S. aureus infections (6,300 infections), while community-acquired infections accounted for 54% (7,250 infections) (Table 6). Community-acquired pneumonia as a primary diagnosis accounted for 12% (1,500) of the total cases. If community-acquired pneumonia is assumed to be mostly acquired in long-term care facilities, most infections (58%) were acquired institutionally. The cost attributable to community-acquired infections ($35,300) was approximately $6,500 higher on a per patient basis than the cost attributable to nosocomial infections ($28,800). The death rates attributable to community-acquired and nosocomial infections were similar (10.5% and 10.1%). Table 6. Incidence, length of stay, costs, and deaths of Staphylococcus aureus infections by source of infection and degree of resistance -------------------------------------------------------------------------- Direct medical cost Per Deaths Total patient Source of infection Incidence ($M) ($) Total (%) -------------------------------------------------------------------------- Nosocomial 6,300 181.0 28,800 640 10 Community 7,250 254.5 5,300 760 11 Pneumonia 1,500 51.7 34,900 380 25 Non-pneumonia 5,750 202.8 35,400 380 7 MRSA(sup a) 2,780 94.5 34,000 590 21 MSSA(sup b) 10,770 339.4 31,500 810 8 -------------------------------------------------------------------------- (sup a)Methicillin-resistant strains of S. aureus. (sup b)Methicillin-sensitive strains of S. aureus. MRSA Infection MRSA infections accounted for 21% (2,780) of the total S. aureus infection incidence (29% of 6,300 nosocomial infections plus 10% of 7,250 community-acquired infections), while MSSA infections accounted for 79% (10,770) of total infections (Table 6). The attributable cost of a patient with MRSA was approximately $2,500 higher than the attributable cost of a patient with MSSA ($34,000 versus $31,500). The higher cost of MRSA infections is due to the higher cost of vancomycin, longer hospital stay, and the cost of patient isolation procedures. For nosocomial infections alone, the cost attributable to MRSA was approximately $3,700 higher on a per patient basis than the cost attributable to MSSA infections ($31,400 versus $27,700). The death rate attributable to MRSA infections was estimated at more than 2.5 times higher than that attributable to MSSA infections (21% versus 8%). Sensitivity Analyses Although assumed to be underused in SPARCS, the ICD-9-CM code 041.11 represents a lower boundary of the total incidence of S. aureus infection. In SPARCS, code 041.11 was used 7,366 times associated with a diagnosis of interest (e.g., endocarditis) and represented a total cost of $236.4 million and a death rate (740 deaths) of 2% (Table 7). The upper boundary of the total cost of S. aureus infections was calculated by assuming that all hospital charges and deaths of patients with S. aureus infections were attributable to the infection, representing a total cost of $599 million and a death rate of 14.5% (1,960 deaths). We conducted sensitivity analyses (varying the percentage of nosocomial MRSA; percentage of patients isolated; difference in length of stay between patients with MRSA and MSSA; attributable length of stay for patients with ventilator- associated pneumonia; number of S. aureus catheter infections; and percentage of S. aureus–caused bacteremia, septicemia, and postoperative infections) and found that the difference in cost per case between MRSA and MSSA infections was $1,700 to $5,100. Table 7. Sensitivity analyses. --------------------------------------------- Direct medical cost ($) Deaths --------------------------------------------- Study results 435.5 1,400 Lower boundary: only 041.11 cases 236.4 740 Upper boundary: all costs attributable 599.0 1,960 --------------------------------------------- Comments Our sensitivity analysis shows that we did not vastly over- or underestimate the direct medical costs of S. aureus infections in New York City. However, the study had several limitations; it was retrospective, and the data sources were not validated by other means (e.g., interviews or chart review). Therefore, coding errors in this database may affect the results. The clinical panel estimates we used to model differences between MRSA and MSSA may lead to some inaccuracy in those difference estimates. Thus, our comparison of costs and deaths between MRSA and MSSA should be viewed as a best approximation in the absence of case-control data or a multivariate analysis of a well-defined patient population. Our estimates of the cost per infection are generally higher than estimates in studies reviewed by Jarvis (19). A major reason may be our focus on New York City, where costs are much higher than in other areas of the United States. In addition, earlier studies have used only hospital costs. Our perspective was societal; therefore, we included physician fees and outpatient costs, as well as hospital charges. Finally, most of these studies focused on nonorganism-specific nosocomial infections; S. aureus infections may have a higher average cost per episode than infections of other organisms (42). On the other hand, we used conservative estimates for certain costs. Medicare prices for professional services are generally lower than commercial rates. Also, we did not account for postdischarge complications that did not lead to hospitalization. Additionally, our societal estimates did not include the cost of dying or lost productivity associated with these illnesses. Despite its limitations, this study shows that hospitalizations associated with S. aureus are serious and have high medical costs and death rates. The average length of stay attributable to S. aureus infection for these patients was very high, 20 days—nearly three times the average for any other type of hospitalization (43). The increased length of stay in turn leads to increases in direct medical costs, with an average cost per case of $32,100 in 1995. Treating an MRSA infection costs 6% to 10% more than treating an MSSA infection $2,500 to $3,700 per case). This cost difference does not reflect MRSA's greater virulence; rather, it reflects the increased cost of vancomycin use and isolation procedures (if used). These estimates are slightly lower than the difference of $5,104 found by Wakefield et al. (21), perhaps because they focused on the cost of serious S. aureus infections, while our analysis examined all hospitalizable S. aureus infections. Patients with MRSA infections have a high average attributable death rate of 21% versus 8% for an MSSA infection. Some of the death rate difference may be related to the underlying condition of patients who become infected with MRSA (e.g., older patients, drug users, sicker patients, patients previously exposed to other antibiotics) (44) and to the lack of effectiveness of vancomycin itself in curing MRSA. (Vancomycin has a narrow therapeutic index that allows little room for increasing blood concentration without incurring substantial losses in tolerance [45]). Both MSSA and MRSA infections are associated with high costs and large numbers of deaths in the New York City metropolitan area. The costs and deaths associated with S. aureus infections may dramatically increase if the newly isolated S. aureus infection with intermediate resistance to vancomycin spreads or if VRSA emerges. For example, after penicillin-resistant S. aureus appeared in the 1950s, the death rate of bacteremia increased from 28% to 50% at the University of Minnesota (Figure) (46). After methicillin was introduced, the death rate decreased (47). Efforts should be directed toward reducing the incidence of MRSA and MSSA nosocomial infections to reduce their economic impact on society. [Fig.] Figure. Death rate of staphylococcal bacteremia over time. (Data from 46, 47.) Acknowledgment We thank the clinical panel: Drs. Donald Armstrong, Donald Low, James Rahal, and Richard B. Roberts. This study was sponsored by the Public Health Research Institute in conjunction with the Bacterial Antibiotic Resistance Group and The Rockefeller University. Funders included The New York Community Trust, The Horace W. Goldsmith Foundation, The United Hospital Fund of New York City, The Texaco Foundation, and the U.S. Centers for Disease Control and Prevention. Dr. Rubin, president of The Lewin Group, a Washington-based health-care consulting company, is a clinical professor of medicine at Georgetown University School of Medicine. From 1981-1984, he was assistant surgeon general in the U.S. Public Health Service and assistant secretary for planning and evaluation, U.S. Department of Health and Human Services. Address for correspondence: Robert J. Rubin, The Lewin Group, 9302 Lee Highway, Fairfax, VA 22031-1214, USA; fax: 703-218-5501. References 1. Haley RW, Culver DH, White JW, Morgan WM, Emori TG. The nationwide nosocomial infection rate: a new need for vital statistics. Am J Epidemiol 1985;121:159. 2. Pittet D, Tarara D, Wenzel RP. Nosocomial bloodstream infection in critically ill patients, excess length of stay, extra costs, and attributable mortality. JAMA 1994;271:1598-601. 3. Brachman PS, Dan BB, Haley RW, Hooten TM, Garner JS, Allen JR. Nosocomial surgical infections: incidence and cost. Surg Clin North Am 1980;60:15-25. 4. Centers for Disease Control and Prevention. National Nosocomial Infection Surveillance System report: data summary from October 1986-April 1996. Atlanta (GA): U.S. Department of Health and Human Services; 1996. 5. Neu HC. The crisis in antibiotic resistance. Science 1992;257:1064-72. 6. Barrett FF, McGehee RF, Finland M. Methicillin-resistant Staphylococcus aureus at Boston City hospital. N Engl J Med 1968;279:441. 7. Boyce JM. Increasing prevalence of methicillin-resistant Staphylococcus aureus in the United States. Infect Control Hosp Epidemiol 1990;11:639-42. 8. Panlilio AL, Culver DH, Gaynes RP, Banerjee S, Henderson TS, Tolson JS, et al. Methicillin-resistant Staphylococcus aureus in U.S. hospitals, 1975-1991. Infect Control Hosp Epidemiol 1992;13:582-6. 9. Nosocomial enterococci resistant to vancomycin—United States, 1989-1993. MMWR Morb Mortal Wkly Rep 1993;42:597-9. 10. Recommendations for preventing the spread of vancomycin resistance recommendations of the Hospital Infection Control Practices Advisory Committee. MMWR Morb Mortal Wkly Rep 1995;44(RR12):1-13. 11. Reduced susceptibility of Staphylococcus aureus to vancomycin—Japan, 1996. MMWR Morb Mortal Wkly Rep 1997;46:624-6. 12. Update: Staphylococcus aureus with reduced susceptibility to vancomycin—United States, 1997. MMWR Morb Mortal Wkly Rep 1997;46:813-5. 13. Boyce JM, Jackson MM, Pugliese G, Batt MD, Fleming D, Garner JS, et al. Methicillin-resistant Staphylococcus aureus (MRSA): a briefing for acute care hospitals and nursing facilities. Infect Control Hosp Epidemiol 1994;15:105-15. 14. McManus AT, Mason AD, McManus WF, Pruitt BA. What's in a name? Is methicillin-resistant Staphylococcus aureus just another S. aureus when treated with vancomycin? Arch Surg 1989;124:1456-9. 15. Pittet D, Waldvogel FA. To control or not to control colonization with MRSA…that's the question! QJM 1997;90:239-41. 16. Teare EL, Barrett SP. Stop the ritual of tracing colonised people. BMJ 1997;314:665-6. 17. Cookson B. Controversies: is it time to stop searching for MRSA? Screening is still important. BMJ 1997;314:664-5. 18. Casewell MW. New threats to the control of methicillin-resistant Staphylococcus aureus. J Hosp Infect 1995;30 Suppl:465-71. 19. Jarvis WR. Selected aspects of the socioeconomic impact of nosocomial infections: morbidity, mortality, cost, and prevention. Infect Control Hosp Epidemiol 1996;17:552-7. 20. Haley RW, White JW, Culver DH, Hughes JM. The financial incentive for hospitals to prevent nosocomial infections under the prospective payment system: an empirical determination from a nationally representative sample. JAMA 1987;257:1611-4. 21. Wakefield DS, Pfaller MA, Hammons GT, Massanari RM. Use of the appropriateness evaluation protocol for estimating the incremental costs associated with nosocomial infections. Med Care 1987;25:481-8. 22. Jernigan JA, Clemence MA, Stott GA, Titus MG, Alexander CH, Palumbo CM, et al. Control of methicillin-resistant Staphylococcus aureus at a university hospital. Infect Control Hosp Epidemiol 1995;16:686-96. 23. Wakefield DS, Helms CM, Massanari RM, Mori M, Pfaller M. Cost of nosocomial infection: relative contributions of laboratory, antibiotic and per diem costs in serious Staphylococcus aureus infections. Am J Infect Control 1988;16:185-92. 24. Frieden TR, Fujiwara PI, Washko RM, Hamburg MA. Tuberculosis in New York City—turning the tide. N Engl J Med 1995;333:229-33. 25. Frieden TR, Munsiff SS, Low DE, Willey BM, Williams G, Faur Y, et al. Emergence of vancomycin-resistant enterococci in New York City. Lancet 1993;342:76-9. 26. New York State Department of Health. 1995 Statewide Planning and Research Cooperative System (SPARCS) Administratively Releasable File. Albany (NY): The Department; 1997. 27. Health Care Financing Administration. Physician fee schedule (CY 1995); payment policies and relative value adjustments. Federal Register 1994;59(235):63410-635. 28. 1995 Drug Topics Red Book. Montvale (NJ): Medical Economics Company; 1995. 29. International classification of diseases, 9th revision, clinical modifier: with color symbols: ICD-9-CM. 4th ed. Salt Lake City (UT): Medicode Publications; 1994. 30. Lautenschlager S, Herzog C, Zimmerli W. Course and outcome of bacteremia due to Staphylococcus aureus: evaluation of different clinical case definitions. Clin Infect Dis 1993;16:567-73. 31. Espersen F. Identifying the patient risk for Staphylococcus aureus bloodstream infections. J Chemother 1995;7:11-7. 32. Muder R, Brennen C, Wagener M, Goetz A. Bacteremia in a long-term care facility: a five year prospective study of 163 consecutive episodes. Clin Infect Dis 1992;14:647-54. 33. Mandell GI, Bennett JE, Dolin R, editors. Mandell, Douglas and Bennett's principles and practices of infectious diseases. 4th ed. New York: Churchill Livingstone; 1995. 34. Lavery LA, Sariaya M, Ashry H, Harkless LB. Microbiology of osteomyelitis in diabetic foot ulcers. J Foot Ankel Surg 1995;34:61-4. 35. Isselbacher KJ, Braunwald E, Wilson JD, Martin JB, Fauci AS, Kasper DL, editors. Harrison's principles of internal medicine. 13th ed. New York: McGraw-Hill, Inc.; 1994. 36. Fagon JY, Chastre J, Vuagnat A, Troillet JL, Novara A, Gibert C. Nosocomial pneumonia and mortality among patients in intensive care units. JAMA 1996;275:866-9. 37. Papazian L, Bregeon F, Thirion X, Gregoire R, Saux P, Denis JP, et al. Effect of ventilator-associated pneumonia on mortality and morbidity. Am J Respir Crit Care Med 1996;154:91-7. 38. Fagon JY, Chastre J, Hance AJ, Montravers P, Novara A, Gibert C. Nosocomial pneumonia in ventilated patients: a cohort study evaluating attributable mortality and hospital stay. Am J Med 1993;94:281-8. 39. Leu HS, Kaiser DL, Mori M, Woolson RF, Wenzel RP. Hospital-acquired pneumonia: attributable mortality and morbidity. Am J Epidemiol 1989;129:1258-67. 40. Craven PC. Treating bone and joint infections with teicoplanin: hospitalization vs. outpatient cost issues. Hospital Formulary 1993;28:41-5. 41. Allen R. Cost-effectiveness issues for home IV therapy in the United States. Hospital Formulary 1993;28:37-40. 42. Arnow PM, Quimosing EM, Beach M. Consequences of intravascular catheter sepsis. Clin Infect Dis 1993;16:778-84. 43. Agency for Health Care Policy and Research. The HCUP-3 Nationwide Inpatient Sample (NIS), Release 2, 1993. Springfield (VA): National Technical Information Service; 1996. 44. Bradley SF. Methicillin-resistant Staphylococcus aureus infection. Clin Geriatr Med 1992;8:853-68. 45. McEvoy GK, editor. American hospital formulary service drug information 1997. Bethesda (MD): American Society of Health-System Pharmacists; 1997. 46. Spink WW. Staphylococcal infections and the problem of antibiotic-resistant staphylococci. Arch Int Med 1954;94:167-196. 47. Allen JD, Roberts CE, Kirby WM. Staphylococcal septicemia treated with methicillin: report of twenty-two cases. N Engl J Med 1962;266:111-6. ——————————————————————————————————————————————————————————————————————————— Perspectives Socioeconomic and Behavioral Factors Leading to Acquired Bacterial Resistance to Antibiotics in Developing Countries Iruka N. Okeke,* Adebayo Lamikanra,* and Robert Edelman† *Obafemi Awolowo University, Ile-Ife, Nigeria; and †University of Maryland School of Medicine, Baltimore, Maryland, USA --------------------------------------------------------------------------- In developing countries, acquired bacterial resistance to antimicrobial agents is common in isolates from healthy persons and from persons with community-acquired infections. Complex socioeconomic and behavioral factors associated with antibiotic resistance, particularly regarding diarrheal and respiratory pathogens, in developing tropical countries, include misuse of antibiotics by health professionals, unskilled practitioners, and laypersons; poor drug quality; unhygienic conditions accounting for spread of resistant bacteria; and inadequate surveillance. Acquired bacterial resistance is common in isolates from healthy persons and from patients with community-acquired infections in developing countries, where the need for antibiotics is driven by the high incidence of infectious disease (1). Among isolates of diarrheal, respiratory, and commensal enteric pathogens (2-5), resistance is increasing, particularly to first-line, inexpensive, broad-spectrum antibiotics (Table 1). Furthermore, introduction of newer drugs (e.g., fluoroquinones) has been followed relatively quickly by the emergence and dissemination of resistant strains (5). The selection and spread of resistant organisms in developing countries, which can often be traced to complex socioeconomic and behavioral antecedents, contribute to the escalating problem of antibiotic resistance worldwide. Table 1. Pathogens with a steadily increasing prevalence of acquired antibiotic resistance in developing tropical countries ----------------------------------------------------------------------------- Pathogen Drug(s) Country (years) Ref. ----------------------------------------------------------------------------- Shigella flexneri, ampicillin, Bangladesh (6) S. dysenteriae tetracycline, (1983-1990) sulfonamides (alone Brazil (1988-1993) (7) of with trimethoprim), Rwanda (1983-1993) (8) nalidixic acid Thailand (1981-1995) (5) Vibrio cholerae cotrimethoxazole, Guinea-Bissau (9) nalidixic acid, (1987-1995) ampicillin India (1993-1995) (10) Salmonella typhi ampicillin, Bangladesh (3) chloramphenicol, (1989-1993) cotrimethoxazole Salmonella cotrimethoxazole Thailand (1981-1995) (5) (nontyphoidal) Enterotoxigenic cotrimethoxazole Thailand (1981-1995) (5) Escherichia coli Campylobacter fluoroquinolones Thailand (1987-1995) (5) Mycobacterium isoniazid, Kenya (1981-1990) (11) tuberculosis streptomycin, Morocco (1992-1994) (12) rifampicin (primary resistance) ----------------------------------------------------------------------------- Misuse of Antibiotics by Physicians in Clinical Practice Antibiotic use provides selective pressure favoring resistant bacterial strains; inappropriate use increases the risk for selection and dissemination of antibiotic-resistant bacteria, which are placed at a competitive advantage. Therefore, one would expect that drugs more commonly affected by bacterial resistance in developing countries are generally inexpensive and popular broad-spectrum agents (2-5,13). However, the relationship between antibiotic use and the emergence and spread of resistance is complex. Antibiotic use in clinical practice alone cannot explain the high frequency of resistant organisms in developing countries (14,15). Nevertheless, excessive clinical use (a form of misuse) is at least partially responsible for the escalating rates of resistance, especially in hospital settings, worldwide. The unnecessary prescription of antibiotics seen in industrialized nations has also been documented in many developing countries, particularly in cases of acute infantile diarrhea and viral respiratory infections (16-22). Clinical misuse of antibiotics may be more common among private practitioners than among public health personnel—private practitioners charge higher fees, the demand for antibiotics seen in private patients is higher, and more drugs are available in private clinics than in public hospitals (23-25). Several strategies have been proposed for combating the inappropriate use of antibiotics by clinicians (26). Antibiotic monitoring systems and hospital formularies or antibiotic treatment protocols often reduce antibiotic prescription rates (24,27). Adoption of a national essential drug list can limit the antibiotics available to prescribers (28,29). However, implementation of these strategies does not guarantee optimal antibiotic use by clinicians in developing countries because the irregular drug supply, availability of drugs from unofficial sources, and financial constraints also affect antibiotic choices (30-32). Continuing medical education changes the attitude of clinicians. Studies of antibiotic misuse in Cuba and Pakistan (33,34) recommend continuing medical education for health workers as the single most important tool for combating antibiotic misuse. A study in Zambia has demonstrated the efficacy of education in reducing antibiotic prescription rates (35). However, education has not been successfully implemented in many developing countries, where too often, governments and health workers cannot afford the time and money required for continuing medical education (36). Health workers in many developing countries have almost no access to objective health information (24). Pharmaceutical company representatives typically outnumber practitioners and often adversely influence their prescription habits (37), as reflected by sales of nonessential drugs and drug combinations (38). Drug labels and package inserts often fail to provide accurate information (39), and in industrialized countries, patients often pressure physicians to prescribe antibiotics (19). Misuse of Antibiotics by Unskilled Practitioners In many developing countries, well-trained health personnel are scarce and cannot serve the entire population, especially in rural areas. Community health workers and others with minimal training treat minor ailments (40). The qualifications and training of community health workers, as well as the quality of care they provide, vary from country to country. Unskilled personnel are less aware of the deleterious effects of inappropriate antibiotic use. For example, pharmacy technicians in Thailand prescribed rifampicin for urethritis and tetracycline for young children (41). Unqualified drug sellers offer alternative drugs when the prescribed drugs are out of stock or refill prescriptions without consulting the prescriber (42,43). In India, traditional healers often dispense antibiotics (44). A high proportion of patients in some developing countries are treated by untrained practitioners simultaneously with oral and injectable antibiotics administered with contaminated needles and syringes (45-47) for misdiagnosed noninfectious diseases (48). Misuse of Antibiotics by the Public In most developing countries, antibiotics can be purchased without prescription, even when the practice is not legal. In many African, Asian, and Latin American countries, antibiotics are readily available on demand from hospitals, pharmacies, patent medicine stalls (drugstores), roadside stalls, and hawkers (17,43,46,49-53). In rural Bangladesh, for example, 95% of drugs consumed for 1 month by more than 2,000 study participants came from local pharmacies; only 8% were prescribed by physicians (54). People are encouraged to buy from unofficial distributors because drugs often are not available in government hospitals (55). Drug vendors usually have little or no knowledge of the required dosage regimen, indications, or contraindications (43,45,55). In markets and public transport in West African countries such as Cameroon (49) and Nigeria (Okeke and Lamikanra, pers. obs.), the vendor (usually a medically untrained salesman) tries to convince potential buyers to purchase the drug, even if they are not ill. To save time and keep drug-hunting to a minimum, a patient may start at a source more likely to stock the desired drug, forgoing the expertise of a doctor. Unofficial sources are generally more accessible than official sources. For example, in Nepal, retail drug outlets are four times as numerous as government health posts and hospitals (46). Alternate sources offer the option of purchasing small quantities of medicines, while hospitals require purchase of the complete 5- or 7-day antibiotic regimen (17,43,52). The purchase of small samples is exceedingly common, particularly for most customers, who buy without prescription (52). These subinhibitory antibiotic regimens predispose for selection of resistant bacterial strains. Antibiotic use in developing countries is underestimated. The quantity of drugs distributed within a country is calculated under the assumption that each person purchases a complete regimen (56). However, medication can be purchased in small aliquots from roadside stalls, and distribution of locally produced or counterfeit antibiotics is not recorded. The motives for self-medication and antibiotic overuse by laypersons are similar to those for clinical abuse by health professionals: to cut costs and act expeditiously to treat confirmed or suspected bacterial infection (57). For example, 50% to 80% of Bangladeshi patients infected with Shigella admitted that they had taken at least one antibiotic in the 15 days before a hospital visit (58), as had 18% to 70% of pediatric patients with acute respiratory infection in two Chinese studies (20,59). The proportion of patients who self-medicate is probably higher, because patients are often reluctant to admit having taken antibiotics before visiting a hospital (60). Common cultural beliefs about antibiotics include the notions that there is a pill for every symptom; antibiotics can heal many illnesses, including dyspepsia and headaches; and injections are more powerful than pills. The misuse of antibiotics frequently becomes integrated into the local culture (62) (e.g., antibiotics are used to prevent diarrhea after eating suspected contaminated foods or [by prostitutes] to prevent sexually transmitted diseases [52,63]). Another cause of antibiotic abuse and selection for resistant bacteria is poor patient compliance. First, physician-patient interactions are often inadequate. They can be short (e.g., a mean of 54 sec was recorded in a Bangladeshi study [16]) and of poor quality (e.g., in Mexico, poor patient-physician communication was partially responsible for the noncompliance of patients with antibiotic regimens [21]). Second, because patients often travel long distances and incur large expenses for medical care, they are unlikely to return for follow-up visits. The reverse situation—the prescriber visiting his patient—is difficult logistically, especially in rural Africa (64). In addition, the patient may be unable to read medicine labels. Finally, because many drugs are expensive, indigent patients purchase incomplete regimens whenever possible and discontinue treatment when symptoms disappear but before the pathogen is eliminated (52). Poor Quality of Antibiotics Lack of Quality Compliance and Monitoring Besides the risk for therapeutic failure, degradation products or adulterants in poor quality antibiotics can produce subinhibitory concentrations in vivo, which increase the selection of resistant strains. Drugs that do not comply with minimum standards are illegal in all countries. However, the quality of many antibiotics and other drugs in developing countries is often below standards in the formulary. In Nigeria for example, substandard ampicillin, ampicillin/cloxacillin, tetracycline, and oxytetracycline capsules have been detected (53,65-67). In many cases, therapeutic failure is the only indication of substandard drugs. Analytic laboratories to detect substandard drugs are uncommon, and when they exist, health workers, distributors, and consumers are often unaware of them. Degraded Antibiotics The shelf lives of drugs developed and marketed in temperate countries are determined by storage temperatures. During distribution in tropical countries, conditions of transport and storage are poorly controlled, and the drugs may be degraded. Ballereau et al. (68) recorded temperatures of 26°C to 40°C and 30% and 90% humidity in Guinea-Bissau during a 2-year period (temperatures of greater than 25°C can degrade antibiotics). Many antibiotics, being heat- and moisture-labile, are particularly vulnerable. Of seven drugs that lost 10% or more of their active constituents when stored in pharmacies in Guinea-Bissau for 2 years, six were antimicrobial drugs (68). Drug consignments are exposed to such adverse conditions during shipment (69) or at tropical ports while they await lengthy port clearance. Drugs are often handled by untrained workers who may store them incorrectly. Hawkers and small traders in Nigeria frequently display large glass jars containing different types of antibiotic capsules mixed together, fully exposed to harsh sunlight and high ambient temperature and humidity. In a Nigerian study of eight batches of tetracycline capsules, only the batch obtained directly from the manufacturer was not excessively degraded and contained active drug levels within formulary limits (Table 2) (53,70). Studies conducted in Thailand and Nigeria demonstrated similar degradation of chloroquine and amoxicillin (67,70). Table 2. Source and quality of tetracycline capsules in a Nigerian suburban town (compiled with data from [53]) ----------------------------------------------------------------- Tetracycline content (% of Content Bioavail- label of ATC(sup a) ability Sample Source claim) (%) (%)(sup b) ----------------------------------------------------------------- C1 Manufacturer(sup c) 105.9 None 100 detected C2 Hospital 107.5 5.3 63.4(sup d) C3 Roadside stall 104.5 1.1 80.5(sup d) C4 Pharmacy 66.1 2.4 65.2(sup d) C5 Patent medicine 84.5 1.9 87.6(sup d) stall C6 Roadside stall 67.8 1.5 Not tested C7 Patent medicine 89.6 1.8 Not stall tested ----------------------------------------------------------------- (sup a)Anhydrotetracycline, one of four tetracycline degradation products. (sup b)Measured from cumulative excretion of tetracycline in the urine of five volunteers. (sup c)Reference standard obtained from the manufacturer. (sup d)Significantly different from C1 (p = 0.01, Wilcoxon signed rank test). Expired Antibiotics Some pharmacologically active drugs produced in countries have expired when distributed in developing countries—they were shipped at the end of the drugs' shelf lives or their clearance and distribution after trancontinental shipment were delayed. Expired drugs may receive new labels, be dumped without a label change, or be donated rather than sold (71-73). Tax deductions and the cost of liquidation are incentives for donating expired or near- expired drugs. Effective enforcement of the World Health Organization (WHO) guidelines on drug donations may curtail such practices (74) Counterfeit Drugs Some drugs sold in developing countries do not contain the concentration of active substances stated on their labels, even at the time of manufacture. These counterfeit drugs flourish, despite efforts of local regulatory agencies to stop their production and distribution (75-77). Approximately 65% of the 751 instances of counterfeit pharmaceuticals reported to WHO or to Interpol from 28 countries in the past 15 years were produced in developing countries (77). Counterfeit drugs include products with little or no active ingredients (e.g., in Nigeria, Indonesia, Brazil, Thailand, Bangladesh, Malaysia, and Francophone African countries [39,76,78,79]) or products for which excipients have been replaced by less expensive alternatives (e.g., substitution of ethylene glycol for propylene glycol in pediatric paracetamol formulations, which caused many deaths in Nigeria, Argentina, Bangladesh, India, and Haiti [76,78]). Counterfeit drugs, like other counterfeit materials, compete favorably in the markets of developing countries. The analytic facilities available to law enforcement agencies often cannot detect these drugs before they reach the patient. Multinational pharmaceutical companies, which probably possess the best analytic facilities for in-house quality assurance in developing countries, try to detect counterfeit drugs to protect their income and reputation; however, such efforts are directed primarily at counterfeits of these companies' own products. Because of the profusion of generic drugs in developing countries, a substantial proportion of counterfeit drugs go undetected. Adulterated Drugs Herbal preparations in developing countries are often adulterated with orthodox medicaments. For example, in one study, 24% of Chinese herbal preparations marketed in Taiwan contained one or more of such adulterants (80). Although the adulteration of such products with antibiotics has not been reported, such practices may be common (81). A Nigerian traditional healer, for example, admitted to 'augmenting' herbal preparations with tetracycline from commercially available capsules (82). Bioinequivalent Antibiotics and Biopharmaceutic Interactions In the last 2 decades, the importance of bioavailability has been underscored by the recognition that chemically equivalent generic drug formulations do not always deliver the expected amount of drug to the bloodstream. Slowly absorbed and acid-labile antibiotics are particularly prone to bioinequivalence and consequent therapeutic failure. In addition, poorly absorbed antibiotics remain in the gut to facilitate the selection of resistant organisms. The few published studies from the developing world have found bioinequivalence in antibiotic formulations, and the problem may be widespread (Table 2) (53,83). Inexpensive generic antibiotics commonly used in developing countries usually are not subject to bioavailability studies. The bioavailability of an antibiotic formulation is modulated by conditions surrounding its administration; conditions unique to developing countries are rarely investigated. Drug combinations used in the tropics but rarely elsewhere may not be optimally absorbed. For example, coadministration of chloroquine and ampicillin lowers the bioavailability of ampicillin (84). A Nigerian meal lowered the biologic availability of orally administered nitrofurantoin (85). Chewing of Khat, a popular Yemeni stimulant, adversely affected the bioavailability of ampicillin and amoxicillin (86). By contrast, the Ayurvedic preparation Trikatu enhanced the absorption of several drugs (87). Whether traditional medicines with antimicrobial properties enhance antibiotic resistance is unknown. Dissemination of Resistant Organisms Crowding and Unhygienic Conditions Residents of developing countries often carry antibiotic-resistant fecal commensal organisms (13,88). Visitors to developing countries passively acquire antibiotic-resistant gut Escherichia coli, even if they are not taking prophylactic antibiotics, which suggests that they encounter a reservoir of antibiotic-resistant strains during travel (89). Apparently healthy people in developing countries carry potentially pathogenic, antibiotic-resistant organisms asymptomatically (90). Several factors, such as urban migration with crowding and improper sewage disposal, encourage the exchange of antibiotic-resistant organisms between people and the exchange of resistance genes among bacteria, thereby increasing the prevalence of resistant strains. In Nigeria, resistant E. coli isolates from persons in an urban metropolis (Lagos) were significantly more likely to be resistant to ampicillin and streptomycin (p < 0.05), and possibly more resistant to sulphathiazole and tetracycline (p < 0.10), than isolates from residents of nearby smaller towns and villages (Table 3) (91). Moreover, strains isolated from Lagos were more likely to show resistance to 4 to 6 of 7 antibiotics tested, whereas strains from rural areas were in most cases resistant to only 0 to 3 antibiotics (91). Table 3. Antibiotic resistance of Escherichia coli strains isolated from residents of an urban area (Lagos) or rural/suburban areas (southwest Nigeria) (from [91]) -------------------------------------------------------- Percentage of resistant isolates Antimicrobial Urban Rural/suburban agent (n = 30) (n = 44) ------------------------------------------------------- Ampicillin(sup a) 53 27 Chloramphenicol 13 14 Streptomycin(sup a) 63 32 Sulphathiazole(sup b) 73 48 Tetracycline(sup b) 87 64 Trimethoprim 53 41 -------------------------------------------------------- (sup a)Significant differences between the two groups at p < 0.05 (Chi-square test). (sup b)Significant differences between the two groups at p < 0.10 (Chi-square test. In 1991, 80% of residents of developing countries had no sanitary facilities for sewage disposal (92). Pipe-borne water, often scarce in developing countries, is not always potable. The development of sanitation and other facilities is not always proportionate to the rapid rises in urban populations (93,94). As urban migration continues, overcrowding increases and hygiene declines, increasing the probability of spread of antibiotic-resistant and commensal pathogens. Potable water, well-ventilated housing and proper waste disposal should reduce infections, the need for antibiotics, and subsequent development of antibiotic resistance. Because tropical conditions encourages the survival of bacteria, more pathogens and commensals are found in tropical environments than in temperate climates (95). The warm and humid tropical climate and the low levels of health care, hygiene, and sanitation contribute to a relatively high prevalence of infectious disease in developing countries. Inadequate Hospital Infection Control Practices Infection control practices in many hospitals in developing countries are rudimentary and often compromised by economic shortfalls and opposing traditional values (96). The resulting nidus of nosocomial pathogens and resistant organisms may be disseminated to the outside community. Improper disposal of hospital waste accentuates such spread. Untreated hospital waste in Uganda was often dumped into public sewers or thrown into rubbish heaps ravaged by scavengers (97). Inadequate Surveillance Susceptibility Testing and Surveillance Information from routine susceptibility testing of bacterial isolates and surveillance of antibiotic resistance, which provides information on resistance trends, including emerging antibiotic resistance, is essential for clinical practice and for rational policies against antibiotic resistance. Bacterial infections are often treated after they become life-threatening, which encourages empirical selection of broad-spectrum antibiotics (98,99). The antibiotic susceptibility pattern of bacterial isolates in much of the developing world is unknown, and little guides empirical prescribing. Susceptibility testing cannot be done readily because equipment, personnel, and consumables are scarce and expensive (59,100). In most all infections, no clinical specimens are cultured. Where available, community-based antibiotic surveillance data may be useful to prescribers in the absence of patient-specific antibiotic-susceptibility results. For example, Ringertz et al. (101) demonstrated that resistance among respiratory pathogens was infrequent in parts of Ethiopia. This information would help local Ethiopian prescribers to treat such infections with inexpensive, broad-spectrum antibiotics. National surveillance programs for antibiotic resistance, the norm in industrialized nations, are less common and less elaborate in developing countries (4). Current inferences about antibiotic resistance trends in developing countries are based on a small number of reports, generated by a handful of microbiology laboratories in urban areas—data not representative of a country, because wide variations in antibiotic resistance patterns may exist within countries (Table 3). Moreover, surveillance should be conducted regularly and continuously because resistance rates can vary in one region of a country over time (Table 1) (102). Defective Antibiotic Susceptibility Assays Well-standardized antibiotic susceptibility assays provide more reliable results (103). However, standard bacterial strains with which to assay new batches of antibiotics or antibiotic disks are not available in laboratories in many developing countries. Delayed transportation and breakdown of cold storage also affects the quality of antibiotics used as diagnostic reagents. Degraded antibiotic powders and antibiotic disks used for susceptibility testing lead to exaggerated estimates of bacterial resistance levels. The frequent recovery of bacteria resistant to the beta-lactams or tetracyclines in tropical countries could reflect, in part, the temperature and moisture lability of test reagents. Laboratory scientists in developing countries face difficulties in obtaining research supplies, which often require them to improvise by, for example, using injectable antibiotic formulations to measure MICs when standard antibiotic powders are not available. The report that clinical microbiologists in developing countries make their own disks from "local blotting papers" (104) illustrates how improvisation can lead to inconsistent laboratory results and unreliable data. Economic and Political Factors Lack of resources hampers implementation of most strategies against antibiotic resistance. Statistics from the World Bank show that developing countries spent $41 per person on health in 1990, compared with the $1,500 per person spent by industrialized countries. Disease prevalence as measured by disability-adjusted life years and by communicable disease in particular is much greater in developing than in industrialized countries (93,105-107). As a result of such gross underfunding, the drug supply is chronically inadequate or at best erratic in health facilities in many countries, including Nigeria (43,105,106). Armed conflicts have recently led to a breakdown in health services and sanitation and rapid dissemination of resistant pathogens, particularly in sub-Saharan Africa and Asia (108,109,110). During an outbreak of cholera and bacillary dysentery in Rwandan refugees, resistance to multiple first-line antibiotics in clinical isolates of Vibrio cholerae and Shigella dysenteriae contributed to high death rates (109). Even in developing countries not at war, political corruption and mismanagement of funds, personnel, and development programs have created large populations living in abject poverty and at high risk for infection (111). Medical expenses, days lost from work, and transportation costs account for substantial economic loss. The cost of medical treatment, even subsidized treatment, is beyond the means of many patients. Poorly paid health workers sometimes extort fees from patients (111). Thus, persons with communicable diseases, unable to afford medical treatment, may infect others. Poverty also interferes with patient compliance, which in turn promotes the emergence of antibiotic resistance during short-term therapy of acute infections and long-term therapy of chronic infections, such as tuberculosis (111). Combating the Problem of Antibiotic Resistance The recommendations of WHO for ensuring proper drug use (79) can be adapted to combat the escalation of community-acquired antibiotic resistance in developing countries. The misuse of antibiotics by health-care professionals, unskilled practitioners, and patients can be alleviated by auditing antibiotics, limiting antibiotic choice, developing prescription guidelines, and emphasizing continuing medical and public education. The quality of antibiotics can be improved by emphasizing quality compliance and monitoring antimicrobial drugs manufactured or dispensed. Such reforms will help control substandard drugs that are degraded, counterfeit, or bioinequivalent. Dissemination of resistant organisms in the community can be impeded by improved public sanitation and hygienic practices and upgraded hospital infection control. Finally, strategies to ensure that these recommendations are adopted and implemented under difficult economic and political conditions can be formulated. Antimicrobial resistance will continue to escalate in developing countries unless corrective measures are instituted. Acknowledgments We thank the International Program in the Chemical Sciences, which supports the current research in antibiotic resistance by I.N. Okeke and A. Lamikanra. Ms. I.N. Okeke thanks the Fulbright Commission and the United States Information Service for a fellowship and Dr. James B. Kaper for his support. 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JAMA 1994;272:377-81. 111. Cornwall J. Tuberculosis: a clinical problem of international importance. Lancet 1997;350:660-1. ——————————————————————————————————————————————————————————————————————————— Perspectives Campylobacter jejuni—An Emerging Foodborne Pathogen Sean F. Altekruse,* Norman J. Stern,† Patricia I. Fields,‡ and David L. Swerdlow‡ *U.S. Food and Drug Administration, Blacksburg, Virginia, USA; †U.S. Department of Agriculture, Athens, Georgia, USA; and ‡Centers for Disease Control and Prevention, Atlanta, Georgia, USA --------------------------------------------------------------------------- Campylobacter jejuni is the most commonly reported bacterial cause of foodborne infection in the United States. Adding to the human and economic costs are chronic sequelae associated with C. jejuni infection—Guillian-Barré syndrome and reactive arthritis. In addition, an increasing proportion of human infections caused by C. jejuni are resistant to antimicrobial therapy. Mishandling of raw poultry and consumption of undercooked poultry are the major risk factors for human campylobacteriosis. Efforts to prevent human illness are needed throughout each link in the food chain. History Awareness of the public health implications of Campylobacter infections has evolved over more than a century (1). In 1886, Escherich observed organisms resembling campylobacters in stool samples of children with diarrhea. In 1913, McFaydean and Stockman identified campylobacters (called related Vibrio) in fetal tissues of aborted sheep (1). In 1957, King described the isolation of related Vibrio from blood samples of children with diarrhea, and in 1972, clinical microbiologists in Belgium first isolated campylobacters from stool samples of patients with diarrhea (1). The development of selective growth media in the 1970s permitted more laboratories to test stool specimens for Campylobacter. Soon Campylobacter spp. were established as common human pathogens. Campylobacter jejuni infections are now the leading cause of bacterial gastroenteritis reported in the United States (2). In 1996, 46% of laboratory-confirmed cases of bacterial gastroenteritis reported in the Centers for Disease Control and Prevention/U.S. Department of Agriculture/Food and Drug Administration Collaborating Sites Foodborne Disease Active Surveillance Network were caused by Campylobacter species. Campylobacteriosis was followed in prevalence by salmonellosis (28%), shigellosis (17%), and Escherichia coli O157 infection (5%) (Figure 1). [Fig] Figure 1. Cases of Campylobacter and other foodborne infections by month of specimen collection; Centers for Disease Control and Prevention/U.S. Department of Agriculture/Food and Drug Administration Collaborating Sites Foodborne Disease Active Surveillance Network, 1996. Disease Prevalence In the United States, an estimated 2.1 to 2.4 million cases of human campylobacteriosis (illnesses ranging from loose stools to dysentery) occur each year (2). Commonly reported symptoms of patients with laboratory- confirmed infections (a small subset of all cases) include diarrhea, fever, and abdominal cramping. In one study, approximately half of the patients with laboratory-confirmed campylobacteriosis reported a history of bloody diarrhea (3). Less frequently, C. jejuni infections produce bacteremia, septic arthritis, and other extraintestinal symptoms (4). The incidence of campylobacteriosis in HIV-infected patients is higher than in the general population. For example, in Los Angeles County between 1983 and 1987, the reported incidence of campylobacteriosis in patients with AIDS was 519 cases per 100,000 population, 39 times higher than the rate in the general population. (5). Common complications of campylobacteriosis in HIV-infected patients are recurrent infection and infection with antimicrobial-resistant strains 6). Deaths from C. jejuni infection are rare and occur primarily in infants, the elderly, and patients with underlying illnesses (2). Sequelae to Infection Guillain-Barré syndrome (GBS), a demyelating disorder resulting in acute neuromuscular paralysis, is a serious sequela of Campylobacter infection (7). An estimated one case of GBS occurs for every 1,000 cases of campylobacteriosis (7). Up to 40% of patients with the syndrome have evidence of recent Campylobacter infection (7). Approximately 20% of patients with GBS are left with some disability, and approximately 5% die despite advances in respiratory care. Campylobacteriosis is also associated with Reiter syndrome, a reactive arthropathy. In approximately 1% of patients with campylobacteriosis, the sterile postinfection process occurs 7 to 10 days after onset of diarrhea (8). Multiple joints can be affected, particularly the knee joint. Pain and incapacitation can last for months or become chronic. Both GBS and Reiter syndrome are thought to be autoimmune responses stimulated by infection. Many patients with Reiter syndrome carry the HLA B27 antigenic marker (8). The pathogenesis of GBS (9) and Reiter syndrome is not completely understood. Treatment of C. jejuni Infections Supportive measures, particularly fluid and electrolyte replacement, are the principal therapies for most patients with campylobacteriosis (10). Severely dehydrated patients should receive rapid volume expansion with intravenous fluids. For most other patients, oral rehydration is indicated. Although Campylobacter infections are usually self limiting, antibiotic therapy may be prudent for patients who have high fever, bloody diarrhea, or more than eight stools in 24 hours; immunosuppressed patients, patients with bloodstream infections, and those whose symptoms worsen or persist for more than 1 week from the time of diagnosis. When indicated, antimicrobial therapy soon after the onset of symptoms can reduce the median duration of illness from approximately 10 days to 5 days. When treatment is delayed (e.g., until C. jejuni infection is confirmed by a medical laboratory), therapy may not be successful (10). Ease of administration, lack of serious toxicity, and high degree of efficacy make erythromycin the drug of choice for C. jejuni infection; however, other antimicrobial agents, particularly the quinolones and newer macrolides including azithromycin, are also used. Antimicrobial Resistance The increasing rate of human infections caused by antimicrobial-resistant strains of C. jejuni makes clinical management of cases of campylobacteriosis more difficult (11,12). Antimicrobial resistance can prolong illness and compromise treatment of patients with bacteremia. The rate of antimicrobial-resistant enteric infections is highest in the developing world, where the use of antimicrobial drugs in humans and animals is relatively unrestricted. A 1994 study found that most clinical isolates of C. jejuni from U.S. troops in Thailand were resistant to ciprofloxacin. Additionally, nearly one third of isolates from U.S. troops located in Hat Yai were resistant to azithromycin (11). In the industrialized world, the emergence of fluoroquinolone-resistant strains of C. jejuni illustrates the need for prudent antimicrobial use in food-animal production (12). Experimental evidence demonstrates that fluoroquinolone-susceptible C. jejuni readily become drug-resistant in chickens when these drugs are administered (13). After flouroquinolone use in poultry was approved in Europe, resistant C. jejuni strains emerged rapidly in humans during the early 1990s (12). Similarly, within 2 years of the 1995 approval of fluoroquinolone use for poultry in the United States, the number of domestically acquired human cases of ciprofloxacin-resistant campylobacteriosis doubled in Minnesota (14). In a 1997 study conducted in Minnesota, 12 (20%) of 60 C. jejuni isolates obtained from chicken purchased in grocery stores were ciprofloxacin-resistant (14). Pathogenesis The pathogenesis of C. jejuni infection involves both host- and pathogen-specific factors. The health and age of the host (2) and C. jejuni-specific humoral immunity from previous exposure (15) influence clinical outcome after infection. In a volunteer study, C. jejuni infection occurred after ingestion of as few as 800 organisms (16). Rates of infection increased with the ingested dose. Rates of illness appeared to increase when inocula were ingested in a suspension buffered to reduce gastric acidity (16). [Fig] Figure 2. Scanning electron microscope image of Campylobacter jejuni, illustrating its corkscrew appearance and bipolar flagella. Source: Virginia-Maryland Regional College of Veterinary Medicine, Blacksburg, Virginia. Many pathogen-specific virulence determinants may contribute to the pathogenesis of C. jejuni infection, but none has a proven role (17). Suspected determinants of pathogenicity include chemotaxis, motility, and flagella, which are required for attachment and colonization of the gut epithelium (Figure 2) (17). Once colonization occurs, other possible virulence determinants are iron acquisition, host cell invasion, toxin production, inflammation and active secretion, and epithelial disruption with leakage of serosal fluid (17). Survival in the Environment Survival of C. jejuni outside the gut is poor, and replication does not occur readily (17). C. jejuni grows best at 37°C to 42°C (18), the approximate body temperature of the chicken (41°C to 42°C). C. jejuni grows best in a low oxygen or microaerophilic environment, such as an atmosphere of 5% O(sub 2), 10% CO(sub 2), and 85% N(sub 2). The organism is sensitive to freezing, drying, acidic conditions (pH < 5.0), and salinity. Sample Collection and Transport If possible, stool specimens should be chilled (not frozen) and submitted to a laboratory within 24 hours of collection. Storing specimens in deep, airtight containers minimizes exposure to oxygen and desiccation. If a specimen cannot be processed within 24 hours or is likely to contain small numbers of organisms, a rectal swab placed in a specimen transport medium (e.g., Cary-Blair) should be used. Individual laboratories can provide guidance on specimen handling procedures (18). Numerous procedures are available for recovering C. jejuni from clinical specimens (18). Direct plating is cost-effective for testing large numbers of specimens; however, testing sensitivity may be reduced. Preenrichment (raising the temperature from 36°C to 42°C over several hours), filtration, or both are used in some laboratories to improve recovery of stressed C. jejuni organisms from specimens (e.g., stored foods or swabs exposed to oxygen) (19). Isolation can be facilitated by using selective media containing antimicrobial agents, oxygen quenching agents, or a low oxygen atmosphere, thus decreasing the number of colonies that must be screened (18,19). Subtyping of Isolates No standard subtyping technique has been established for C. jejuni. Soon after the organism was described, two serologic methods were developed, the heat-stable or somatic O antigen (20) and the heat-labile antigen schemes (21). These typing schemes are labor intensive, and their use is limited almost exclusively to reference laboratories. Many different DNA-based subtyping schemes have been developed, including pulsed-field gel electrophoresis (PFGE) and randomly amplified polymorphic DNA (RAPD) analysis (22). Various typing schemes have been developed on the basis of the sequence of flaA, encoding flagellin (23); however, recent evidence suggests that this locus may not be representative of the entire genome (24). Transmission to Humans Most cases of human campylobacteriosis are sporadic. Outbreaks have different epidemiologic characteristics from sporadic infections (2). Many outbreaks occur during the spring and autumn (2). Consumption of raw milk was implicated as the source of infection in 30 of the 80 outbreaks of human campylobacteriosis reported to CDC between 1973 and 1992. Outbreaks caused by drinking raw milk often involve farm visits (e.g., school field trips) during the temperate seasons. In contrast, sporadic Campylobacter isolates peak during the summer months (Figure 1). A series of case-control studies identified some risk factors for sporadic campylobacteriosis, particularly handling raw poultry (25,26) and eating undercooked poultry (27-31) (Table). Other risk factors accounting for a smaller proportion of sporadic illnesses include drinking untreated water (29); traveling abroad (25); eating barbequed pork (28) or sausage (27); drinking raw milk (29,32) or milk from bird-pecked bottles (33); and contact with dogs (27) and cats (29,31), particularly juvenile pets or pets with diarrhea (25,34). Person-to-person transmission is uncommon (25,32). Overlap is reported between serotypes of C. jejuni found in humans, poultry, and cattle, indicating that foods of animal origin may play a major role in transmitting C. jejuni to humans (35). In the United States, infants have the highest age-specific Campylobacter isolation rate, approximately 14 per 100,000 person years. As children get older, isolation rates decline to approximately 4 per 100,000 person years for young adolescents. A notable feature of the epidemiology of human campylobacteriosis is the high isolation rate among young adults, approximately 8 per 100,000 person years. Among middle-aged and older adults, the isolation rate is < 3 per 100,000 person years (2). The peak isolation rate in neonates and infants is attributed in part to susceptibility on first exposure and to the low threshold for seeking medical care for infants (2). The high rate of infection during early adulthood, which is pronounced among men, is thought to reflect poor food-handling practices in a population that, until recently, relied on others to prepare meals (2). Reservoirs The ecology of C. jejuni involves wildlife reservoirs, particularly wild birds. Species that carry C. jejuni include migratory birds—cranes, ducks, geese (36), and seagulls (37). The organism is also found in other wild and domestic bird species, as well as in rodents (38). Insects can carry the organism on their exoskeleton (39). Table. Epidemiologic studies of laboratory-confirmed cases of sporadic campylobacteriosis -------------------------------------------------------------------------- Foods Number associated with Animal Cases Controls Date Population Location illness contacts Ref. -------------------------------------------------------------------------- 52 103 1989- Residents Norway Poultry, Dogs 27 1990 of three sausage counties 218 526 1982- HMO Washington Undercooked Animals 30, 1983 patients State chicken with 34 diarrhea 29 42 1990 Residents England Bottled 33 of milk(sup a) Manchester 45 45 1983- University Georgia Chicken Cats 31 1984 students 53 106 1982- Rural Iowa Raw milk 32 1983 children 40 80 1981 Residents Colorado Untreated Cats 29 of Denver, water, Ft. Collins raw milk, undercooked chicken 54 54 1982 Residents Netherlands Chicken, 28 of pork, Rotterdam barbequed foods 10 15 1982 Residents Colorado Preparing 26 of chicken Larimer County 55 14 1980 Residents Sweden Preparing Kitten, 25 of chicken dog Göteborg with diarrhea -------------------------------------------------------------------------- (sup a)Bottle tops pecked by wild birds. The intestines of poultry are easily colonized with C. jejuni. Day-old chicks can be colonized with as few as 35 organisms (40). Most chickens in commercial operations are colonized by 4 weeks (41,42). Vertical transmission (i.e., from breeder flocks to progeny) has been suggested in one study but is not widely accepted (43). Reservoirs in the poultry environment include beetles (39), unchlorinated drinking water (44), and farm workers (41,42,45). Feeds are an unlikely source of campylobacters since they are dry and campylobacters are sensitive to drying. C. jejuni is a commensal organism of the intestinal tract of cattle (46). Young animals are more often colonized than older animals, and feedlot cattle are more likely than grazing animals to carry campylobacters (47). In one study, colonization of dairy herds was associated with drinking unchlorinated water (48). Campylobacters are found in natural water sources throughout the year. The presence of campylobacters is not clearly correlated with indicator organisms for fecal contamination (e.g., E. coli)(49). In temperate regions, organism recovery rates are highest during the cold season (49,50). Survival in cold water is important in the life cycle of campylobacters. In one study, serotypes found in water were similar to those found in humans (50). When stressed, campylobacters enter a "viable but nonculturable state," characterized by uptake of amino acids and maintenance of an intact outer membrane but inability to grow on selective media; such organisms, however, can be transmitted to animals (51). Additionally, unchlorinated drinking water can introduce campylobacters into the farm environment (44,48). Campylobacter in the Food Supply C. jejuni is found in many foods of animal origin. Surveys of raw agricultural products support epidemiologic evidence implicating poultry, meat, and raw milk as sources of human infection. Most retail chicken is contaminated with C. jejuni; one study reported an isolation rate of 98% for retail chicken meat (52). C. jejuni counts often exceed 10(sup 3) per 100 g. Skin and giblets have particularly high levels of contamination. In one study, 12% of raw milk samples from dairy farms in eastern Tennessee were contaminated with C. jejuni (53). Raw milk is presumed to be contaminated by bovine feces; however, direct contamination of milk as a consequence of mastitis also occurs (54). Campylobacters are also found in red meat. In one study, C. jejuni was present in 5% of raw ground beef and in 40% of veal specimens (55). Control of Campylobacter Infection On the Farm Control of Campylobacter contamination on the farm may reduce contamination of carcasses, poultry, and red meat products at the retail level (27). Epidemiologic studies indicate that strict hygiene reduces intestinal carriage in food-producing animals (41,42,45). In field studies, poultry flocks that drank chlorinated water had lower intestinal colonization rates than poultry that drank unchlorinated water (42,44). Experimentally, treatment of chicks with commensal bacteria (56) and immunization of older birds (57) reduced C. jejuni colonization. Because intestinal colonization with campylobacters readily occurs in poultry flocks, even strict measures may not eliminate intestinal carriage by food-producing animals (39,41). At Processing Slaughter and processing provide opportunities for reducing C. jejuni counts on food-animal carcasses. Bacterial counts on carcasses can increase during slaughter and processing steps. In one study, up to a 1,000-fold increase in bacterial counts on carcasses was reported during transportation to slaughter (58). In studies of chickens (59) and turkeys (60) at slaughter, bacterial counts increased by approximately 10- to 100-fold during defeathering and reached the highest level after evisceration. However, bacterial counts on carcasses decline during other slaughter and processing steps. In one study, forced-air chilling of swine carcasses caused a 100-fold reduction in carcass contamination (61). In Texas turkey plants, scalding reduced carcass counts to near or below detectable levels (60). Adding sodium chloride or trisodium phosphate to the chiller water in the presence of an electrical current reduced C. jejuni contamination of chiller water by 2 log(sub 10) units (62). In a slaughter plant in England, use of chlorinated sprays and maintenance of clean working surfaces resulted in a 10- to 100-fold decrease in carcass contamination (63). In another study, lactic acid spraying of swine carcasses reduced counts by at least 50% to often undetectable levels (64). A radiation dose of 2.5 KGy reduced C. jejuni levels on retail poultry by 10 log(sub 10) units (65). Conclusions C. jejuni, first identified as a human diarrheal pathogen in 1973, is the most frequently diagnosed bacterial cause of human gastroenteritis in the United States. Sequelae including GBS and reactive arthritis are increasingly recognized, adding to the human and economic cost of illness from human campylobacteriosis. The emergence of fluoroquinolone-resistant infections in Europe and the United States, temporally associated with the approval of fluoroquinolone use in veterinary medicine, is also a public health concern. The consumption of undercooked poultry and cross-contamination of other foods with drippings from raw poultry are leading risk factors for human campylobacteriosis. Reinforcing hygienic practices at each link in the food chain—from producer to consumer—is critical in preventing the disease. Dr. Altekruse is a Public Health Service Epidemiology Fellow with the Food and Drug Administration, Center for Veterinary Medicine. His current research interest is antimicrobial-resistant foodborne pathogens. Address for correspondence: Sean Altekruse, Virginia-Maryland Regional College of Veterinary Medicine, Duckpond Road, Blacksburg, VA, 24060, USA; fax: 540-231-7367; e-mail: saltekru@vt.edu. References 1. Kist M. The historical background of Campylobacter infection: new aspects. 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RAPD analysis of environmental, food and clinical isolates of Campylobacter spp. FEMS Immunol Med Microbiol 1997;18:119-24. 23. Meinersmann RJ, Helsel LO, Fields PI, Hiett KL. Discrimination of Campylobacter jejuni isolates by fla gene sequencing. J Clin Microbiol 1997;35:2810-4. 24. Harrington CS, Thomson-Carter FM, Carter PE. Evidence for recombination in the flagellin locus of Campylobacter jejuni: implications for the flagellin gene typing scheme. J Clin Microbiol 1997;35:2386-92. 25. Norkrans G, Svedhem Å. Epidemiologic aspects of Campylobacter jejuni enteritis. Journal of Hygiene (Cambridge) 1982;89:163-70. 26. Hopkins RS, Scott AS. Handling raw chicken as a source for sporadic Campylobacter jejuni infections [letter]. J Infect Dis 1983;148:770. 27. Kapperud G, Skjerve E, Bean NH, Ostroff SM, Lassen J. Risk factors for sporadic Campylobacter infections: results of a case-control study in southeastern Norway. J Clin Microbiol 1992;30:3117-21. 28. Oosterom J, den Uyl CH, Bänffer JRJ, Huisman J. Epidemiologic investigations on Campylobacter jejuni in households with primary infection. Journal of Hygiene (Cambridge) 1984;92:325-32. 29. Hopkins RS, Olmsted R, Istre GR. Endemic Campylobacter jejuni infection in Colorado: identified risk factors. Am J Public Health 1984;74:249-50. 30. Harris NV, Weiss NS, Nolan CM. The role of poultry and meats in the etiology of Campylobacter jejuni/coli enteritis. Am J Public Health 1986;76:407-11. 31. Deming MS, Tauxe RV, Blake PA. Campylobacter enteritis at a university from eating chickens and from cats. Am J Epidemiol 1987;126:526-34. 32. Schmid GP, Schaefer RE, Plikaytis BD, Schaefer JR, Bryner JH, Wintermeyer LA, et al. A one-year study of endemic campylobacteriosis in a midwestern city: association with consumption of raw milk. J Infect Dis 1987;156:218-22. 33. Lighton LL, Kaczmarski EB, Jones DM. A study of risk factors for Campylobacter infection in spring. 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Journal of Applied Bacteriology 1992;73:279-85. 39. Jacobs-Reitsma WF, van de Giessen AW, Bolder NM, Mulder RWAW. Epidemiology of Campylobacter spp. at two Dutch broiler farms. Epidemiol Infect 1995;114:413-21. 40. Kaino K, Hayashidani H, Kaneko K, Ogawa M. Intestinal colonization of Campylobacter jejuni in chickens. Japanese Journal of Veterinary Science1988;50:489-94. 41. Humphrey TJ, Henley A, Lanning DG. The colonization of broiler chickens with Campylobacter jejuni; some epidemiologic investigations. Epidemiol Infect 1993;110:601-7. 42. Kapperud G, Skjerve E, Vik L, Hauge K, Lysaker A, Aalmen I, et al. Epidemiological investigation of risk factors for Campylobacter colonization in Norwegian broiler flocks. Epidemiol Infect 1993;111:45-55. 43. Pearson AD, Greenwood MH, Feltham RK, Healing TD, Donaldson J, Jones DM, et al. Microbial ecology of Campylobacter jejuni in a United Kingdom chicken supply chain: intermittent common source, vertical transmission, and amplification by flock propagation. Appl Environ Microbiol 1996;62:4614-20. 44. Pearson AD, Greenwood M, Healing TD, Rollins D, Shahamat M, Donaldson J, et al. Colonization of broiler chickens by waterborne Campylobacter jejuni. Appl Environ Microbiol 1993;59:987-96. 45. Kazwala RR, Collins JD, Hannan J, Crinion RAP, O'Mahony H. Factors responsible for the introduction and spread of Campylobacter jejuni infection in commercial poultry production. Vet Rec 1990;126:305-6. 46. Fricker CR, Park RWA. A two year study of the distribution of thermophilic campylobacters in human, environmental and food samples from the Reading area with particular reference to toxin production and heat stable serotype. Journal of Applied Bacteriology 1989;66:477-90. 47. Giacoboni GI, Itoh K, Hirayama K, Takahashi E, Mitsuoka T. 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Prevalence of Listeria monocytogenes, Campylobacter jejuni, Yersinia enterocolitica, and Salmonella in bulk tank milk: risk factors and risk of human exposure. Journal of Food Protection 1992;55:93-7. 54. Hudson PJ, Vogt RL, Brondum J, Patton CM. Isolation of Campylobacter jejuni from milk during an outbreak of campylobacteriosis. J Infect Dis 1984;150:789. 55. Lammerding AM, Garcia MM, Mann ED, Robinson Y, Dorward WJ, Truscott RB, et al. Prevalence of Salmonella and thermophilic Campylobacter in fresh pork, beef, veal, and poultry in Canada. Journal of Food Protection 1988;51:47-52. 56. Stern NJ. Mucosal competitive exclusion to diminish colonization of chickens by Campylobacter jejuni. Poult Sci 1994;73:402-7. 57. Widders PR, Perry R, Muir WI, Husband AJ, Long KA. Immunization of chickens to reduce intestinal colonization with Campylobacter jejuni. Br Poult Sci 1996;37:765-8. 58. Stern NJ, Clavero MRS, Bailey JS, Cox NA, Robach MC. Campylobacter spp. in broilers on the farm and after transport. Poult Sci 1995;74:937-41. 59. Izat AL, Gardner FA, Denton JH, Golan FA. Incidence and levels of Campylobacter jejuni in broiler processing. Poult Sci 1988;67:1568-72. 60. Acuff GR, Vanderzant C, Hanna MO, Ehlers JG, Golan FA, Gardner FA. Prevalence of Campylobacter jejuni in turkey carcasses during further processing of turkey products. Journal of Food Protection 1986;49:712-7. 61. Oosterom J, De Wilde GJA, De Boer E, De Blaauw LH, Karman H. Survival of Campylobacter jejuni during poultry processing and pig slaughtering. Journal of Food Protection 1983;46:702-6. 62. Li YB, Walker JT, Slavik MF, Wang H. Electrical treatment of poultry chiller water to destroy Campylobacter jejuni. Journal of Food Protection 1995;58:1330-4. 63. Mead GC, Hudson WR, Hinton MH. Effect of changes in processing to improve hygiene control on contamination of poultry carcasses with Campylobacter. Epidemiol Infect 1995;115:495-500. 64. Epling LK, Carpenter JA, Blankenship LC. Prevalence of Campylobacter spp. and Salmonella spp. on pork carcasses and the reduction effected by spraying with lactic acid. Journal of Food Protection 1993;56:536-7,540. 65. Patterson MF. Sensitivity of Campylobacter spp. to irradiation in poultry meat. Letters in Applied Microbiology 1995;20:338-40. —————————————————————————————————————————————————————————————————————————— Synopses —————————————————————————————————————————————————————————————————————————— Synopses Comparative Genomics and Host Resistance against Infectious Diseases Salman T. Qureshi,*† Emil Skamene*† and Danielle Malo*† *McGill University, Montréal, Canada; †Montréal General Hospital, Montréal,Canada --------------------------------------------------------------------------- The large size and complexity of the human genome have limited the identification and functional characterization of components of the innate immune system that play a critical role in front-line defense against invading microorganisms. However, advances in genome analysis (including the development of comprehensive sets of informative genetic markers, improved physical mapping methods, and novel techniques for transcript identification) have reduced the obstacles to discovery of novel host resistance genes. Study of the genomic organization and content of widely divergent vertebrate species has shown a remarkable degree of evolutionary conservation and enables meaningful cross-species comparison and analysis of newly discovered genes. Application of comparative genomics to host resistance will rapidly expand our understanding of human immune defense by facilitating the translation of knowledge acquired through the study of model organisms. We review the rationale and resources for comparative genomic analysis and describe three examples of host resistance genes successfully identified by this approach. Two major elements underlie a thorough understanding of the pathogenesis of virtually any infectious disease: identification and characterization of the virulence factors and in vivo survival mechanisms of the invading microorganism (e.g., surface attachment factors, exotoxins, or enzymes that disrupt cellular homeostasis [1]) and understanding of the components of the host response that lead to elimination of the invading pathogen and resolution of disease. (These include both nonspecific [or innate] immune defense mechanisms, such as the complement cascade, and adaptive elements, such as clonally derived lymphocytes capable of eliminating specific targets [2]). The traditional approach to human infectious diseases has been to focus research on the study of important pathogens. The outcome of investigation of relevant bacteria, viruses, fungi, and parasites has led to the production of protective vaccines, antimicrobial agents, and effective strategies for control and elimination of disease outbreaks. A principal advantage of microbiologic research is the relative ease with which the organisms may be obtained, manipulated, and analyzed in the laboratory. Because microbial genomes are smaller, complete cloning and DNA sequencing of several microorganisms have been achieved and have paved the way for comprehensive study of gene expression and genome organization (3,4). In contrast are relatively limited advances in our understanding of the molecular basis of host defense. The study of host immune defense in humans is inherently complex; obstacles to greater understanding include limited opportunities for controlled observation and experimental manipulation, a large genome, and until recently, a lack of molecular techniques capable of facilitating genomewide analysis. Genetic Analysis One of the principal aims of the study of host response to infectious diseases is to uncover novel components of the host immune system critical to robust host defense. Identification of these components at a molecular level is the first step in understanding how the host deals with an infectious challenge and lays a foundation upon which rational therapies that augment host resistance may someday be designed. Despite this promise, the interaction between host and pathogen that leads to infection is multidimensional, dynamic, and exceedingly complex. From a genomic perspective, a thorough understanding of the pathogenesis of a given infection would include a complete inventory of the spatial and temporal expression of the genes by both the host and pathogen from the time of exposure to the final resolution of the infection. Given the potentially large number of factors that contribute to host defense, precise gene identification is a formidable challenge. Nevertheless, researchers have recently made progress in dissecting and identifying the most important individual genetic elements that govern the host response to important pathogens——largely through the use of animal models of human disease (5). Of the model organisms amenable to genetic analysis, the mouse is by far the most well-developed and physiologically relevant system for study of human host defense (6,7). Identification of commercially available inbred strains of mice that show a differential response to a well-defined infectious challenge is the first requirement for study of genetically regulated host resistance factors. Once distinct phenotypes are identified, controlled breeding is carried out to determine the mode of inheritance of the phenotype (simple or complex). Correlation of the inheritance of susceptibility or resistance to a specific infectious challenge with one or more chromosomal regions is then performed by using linkage analysis. Finally, known genes within the genetic interval must be evaluated and novel genes must be positionally cloned to elucidate the underlying molecular basis of immune defense. Comparative genomic analysis is a logical extension of these principles (8). Knowledge of the genomic organization of human and mouse, for example, facilitates direct localization and identification of the human orthologues of susceptibility genes identified through experimental challenge. These genes can then be tested as candidates for human disease susceptibility through mutation analysis. Genetic Linkage Maps Genetic linkage maps provide an organizational framework for genes and phenotypes in the genome (9). Maps, by establishing the location, order, and relative distance of genes, anonymous DNA markers, and biologically important traits along a species' chromosomes, are critical tools in analyzing genetic contribution to a given disease state. Genetic maps can help precisely localize chromosomal region(s) linked to host resistance phenotypes and provide the starting point for identification of the causative gene(s). During the past decade, comprehensive genetic maps spanning the genomes of mouse and human, have been created largely through the initiative of the Human Genome Project (10). Mapping the Human Genome A great deal of effort has resulted in the creation of a whole genome human linkage map, consisting of 5,624 microsatellite markers located to 2,335 positions (11). The DNA markers in this map are highly informative and are densely distributed, with an average interval between markers of 1.6 centimorgans (cM) (1 cM = a 1% rate of recombination during meiosis, or approximately 1 million bp). Other comprehensive maps have been assembled on the basis of a collection of more than 16,000 distinct transcribed sequences (including known genes and gene fragments) or expressed sequence tags, which are estimated to represent at least 50% of all genes in the human genome (12). This human transcription map has been integrated with selected microsatellite markers from the Généthon collection, thus allowing the position of gene-based markers to be resolved to specific intervals measured in centimorgans. The map is available electronically (13). Work is also under way to generate comprehensive physical maps of the human genome in which the relative location of markers is defined by the actual length along the chromosome, rather than by recombination events (14,15). Mapping the Mouse Genome Among model organisms, genetic mapping is most well established in the mouse, having begun in 1915 with the discovery of the first linkage group (16). Controlled crosses of common laboratory strains segregating a small number of visible phenotypes such as coat color then became the mainstay of genetic mapping. In the past decade, two major breakthroughs have revolutionized the technique of mouse genetic mapping and paved the way for generation of high-resolution whole genome maps. The first was the development of the interspecific cross, involving a laboratory strain (Mus musculus) and a distantly related species Mus spretus (17), allowing literally thousands of genes to be mapped within the same cross. The second advance was the development of abundant genetic markers rapidly typable by polymerase chain reaction (PCR) (termed microsatellites), which amplified polymorphisms in simple sequence length repeats such as [CA]n (Figure) (18). Several comprehensive genetic maps of the mouse (based on genes or microsatellites) have been developed, and in some cases, these are being integrated. At least three are publicly available, while the others are available for mapping in a collaborative arrangement (19). As of January 1997, more than 17,000 markers had been mapped in the mouse (one locus approximately every 200kb), including more than 5,000 genes and more than 10,000 (mostly microsatellite) DNA markers. [Fig] Figure. Schematic representation of microsatellite marker analysis in mice. A) Flanking forward (F) and reverse (R) oligonucleotides are designed to specifically amplify a simple sequence repeat by polymerase chain reaction (PCR) (in this case a CA dinucleotide). The length of the dinucleotide (N) varies among inbred mouse strains. B) Gel electrophoresis of a PCR-amplified microsatellite in homozygous parental strains A and B and heterozygous F1 progeny. The larger microsatellite from strain A migrates more slowly than that of strain B. Inheritance of both parental alleles is shown in the F1. Mapping in Other Species Genetic mapping has been widely embraced by the scientific community; more than 30 vertebrate species are the subject of genetic mapping projects, and high-resolution maps of microsatellite markers have been developed for humans, mice, rats, cows, sheep, pigs, fish, and chickens (19). Two invertebrates, Drosophila melanogaster (a dipteran fly) and Caenorhabditis elegans (a nematode), also have complete genetic and physical maps; the complete nucleotide sequence of the latter is expected in the near future. The status of individual genetic mapping projects and resources has been summarized, along with a compilation of databases for species-specific or comparative mapping reference (19,20). Integrating the data from these species- specific projects in a form that allows relevant information from diverse organisms to be assembled is a major challenge to biologic information systems. The most extensive coverage of mammalian species homologies is the Mouse Genome Database of The Jackson Laboratory (21). Initially developed for the mouse, comparative mapping data for more than 55 species may be searched online, with links to related genomic resources, such as the Human Genome Database, Ratmap, SheepBase, and PigBase. Comparative Genetic Mapping Because of the density of genetic markers positioned along the chromosomes of both organisms, the comparative map of the mouse and human genomes is the most well developed of all species. In a comprehensive summary of mouse/human homology published in 1996, 1,416 loci were placed on both maps by using human physical mapping data and mouse genetic maps (22). This comparison defined 181 conserved linkage groups, approximately 90% of the mouse genome. Further comparative mapping with newly discovered genes and expressed sequence tags will refine the chromosomal relationships between mouse and human. Integrating Maps and Aligning Genomes The integration of existing genetic maps of different species is a formidable challenge. Accurate, comprehensive comparisons of gene arrangements across different species will rapidly advance our understanding of all aspects of biology by allowing rapid information exchange across different model organisms and experimental systems. Several approaches have been used in developing universal mapping probes for diverse genomes (23-25). Of the two classes of loci used to construct gene maps, coding gene sequences (Type I markers), which show conservation among distantly related mammalian species, are most useful as landmarks for comparing linkage and syntenic association. Highly polymorphic sequences (Type II markers), such as microsatellites, are more abundant and are invaluable for mapping within a pedigree but are less useful for comparative purposes because they do not show adequate sequence conservation to recognize locus homology between mammalian orders. In 1993, a list of anchored reference loci for comparative genome mapping in mammals was proposed; it consists of 321 Type I markers equivalently spaced throughout the mammalian genome (26). This approach allowed the position of homologous loci in the maps of four species (human, mouse, cattle, and cat), which represent different mammalian orders, to be established. Interspecies comparison of conserved exon sequences of homologous genes has generated a new overlapping set of anchor loci called comparative anchor tagged sequences (25). Large-scale mapping of these sequences in several species may be an efficient way of developing high-resolution comparative maps with essentially complete genome coverage. Alternative Techniques for Comparative Genomic Analysis Mammalian genomes may be compared at several levels by using a variety of tools and strategies tailored to individual objectives. Although direct sequence comparison of whole genomes will provide the highest resolution for comparative study, this sophisticated form of analysis is at least several years away from being realized. At a cytologic level, species may be compared by fluorescence in situ hybridization (FISH) with single or multiple probes (single or multicolor Zoo-FISH), producing rapid, high-resolution chromosomal localization detectable by microscopy. Alternatively, libraries from microdissected or individual flow-sorted chromosomes may be constructed and used as fluorescence-labeled chromosome "paints" to probe the chromosomes of other species and identify homologous regions (27,28). The main advantage of chromosome painting is its rapid overall evaluation of the extent and character of genomic conservation among distantly related species, such as pig and cattle. In contrast to FISH, chromosome painting does not allow determination of gene order or high-resolution demarcation of chromosomal breakpoints. Radiation hybrid panels, another method for physical assignment of homologous loci (29,30), are generated by irradiation and subsequent fusion of a cell line containing a chromosome from one species, such as human, on another background, such as hamster. The donor DNA is fragmented at random, resulting in a series of lines retaining only fragments of the original chromosome. Conserved genes from other species may be mapped to the homologous region of the human genome by comparing the PCR pattern for each cell line to reference loci with well-established map positions. Models of Human Disease Identifying genetically regulated host immune responses might significantly advance our understanding of the molecular targets and immunologic mechanisms critical to robust defense against pathogenic microbes. To date the number of host defense genes that have been cloned remains small; comparative genomics has the potential to accelerate gene discovery by allowing available data for model organisms to be rapidly applied to the study of human disease. We summarize three examples of human host resistance genes in the following section; in each example, genetic analysis of mouse models of the human disease phenotype played a crucial role in the initial discovery of the human homologue or served as a means of validating the identity of the proposed human candidate disease gene. Nramp 1 and NRAMP1 The Mouse Nramp1 Gene In classic inbred strains of mice, natural resistance to infection with Mycobacterium bovis (BCG), M. lepraemurium, Salmonella Typhimurium, and Leishmania donovani is controlled by the Bcg locus, also known as Ity and Lsh (31-33). The major effect of the Bcg gene is to modulate the growth rate of these diverse pathogens in cells of the reticuloendothelial system of the mouse during the preimmune phase of the infection (33). Resistant and susceptible strains are distinguished by the kinetics of infection shown by pathogen counts (CFUs or Leishmania-forming units) in liver and spleen after infection. The susceptible phenotype is characterized by a higher net growth rate of BCG, Salmonella, or Leishmania in the reticuloendothelial system during the early phase of infection, followed by specific immune responses in BCG- and L. donovaniinfected mice or by a rapidly lethal infection with the virulent pathogen S. Typhimurium. Bcg is inherited as a simple autosomal dominant Mendelian trait in crosses between classical strains of laboratory mice; it was localized to mouse chromosome 1 by linkage analysis (34). Using a positional cloning strategy, Vidal et al. (35) isolated the Nramp1 (natural resistance-associated macrophage protein 1) gene as a strong candidate for the Bcg mutation based on its map location, its macrophage-restricted expression pattern and a nonconservative Gly(sup 169)Asp substitution in the protein of all susceptible strains. Creation of a null allele at Nramp1 then provided formal proof that a mutation within Nramp1 is the cause of the mouse susceptibility to infection with M. bovis, S. Typhimurium, and L. donovani (36). Nramp1, an integral membrane phosphoglycoprotein located in the late endosome/lysosome compartment of resting macrophages, is recruited to the maturing phagosomal membrane (37), consistent with its potential function in controlling the replication of intracellular parasites by altering the intravacuolar environment in which they reside. Nramp1 is part of an ancient family of proteins with highly conserved members in mammals (including humans, cows, rats, sheep), birds, invertebrates (C. elegans, D. melanogaster), plants (Oryza sativa, Arabidopsis thaliana), fungi (Saccharomyces cerevisiae), and even bacteria (M. leprae and Escherichia coli) (38,39). This family is characterized by a highly conserved hydrophobic core consisting of 10 transmembrane (TM) domains with a structural organization typical of families of ion transporters and channels. In addition, the most highly conserved segments of the Nramp family (TM8-TM9 intracellular loop) show impressive similarity with the highly conserved region of mammalian voltage-gated K(sup +) channels of the shaker type (40). Several issues concerning the biochemical function of Nramp1 with respect to intracellular survival of taxonomically unrelated pathogens remain unresolved. Studies of the function of Nramp1-related sequences (Nramp2 and Smf1 in model organisms) provide insight into how Nramp1 confers resistance to microbial agents. Nramp2 has been isolated in mouse and human and shows a high degree of similarity to Nramp1 (77% overall similarity), with identical hydropathy profiles and predicted secondary structures (41,42). Mouse and human Nramp2 mRNA are both widely expressed in contrast with the tissue-specific expression of Nramp1 (41,42). Recently, Nramp2 was shown to be a metal ion transporter with broad divalent cation specificity (including Fe(sup 2+), Zn(sup 2+), Mn(sup 2+), Co(sup 2+), Cd(sup 2+), Cu(sup 2+), Ni(sup 2+), and Pb(sup 2+)), driven by the proton electrochemical gradient in Xenopus laevis oocytes (43). Studies using the yeast double mutant SMF1/ SMF2 provided additional support concerning the function of Nramp2 as a divalent cation transporter. Inactivation of SMF1 and SMF2, two yeast Nramp homologues encoding divalent cation transporters (44), is specifically complemented by Nramp2 (45). In vivo, Nramp2 plays an important role in normal iron transport. Mutation within Nramp2 causes microcytic anemia in mk mutant mice because of severe defects in intestinal iron uptake (46). Interestingly, the missense mutations in mutant Nramp1 and Nramp2 alleles introduce a charged amino acid in two adjacent positions of TM4, confirming the importance of this region of both proteins for normal function. It has been suggested that Nramp1 may also be a divalent cation transporter; its role in reticuloendothelial cells remains unexplored (40,44). The Chicken NRAMP1 Gene The discovery of Nramp1 allowed the study of its role in susceptibility to related infections in other species. Salmonellosis, one of the most common causes of food poisoning in humans, is frequently caused by ingestion of contaminated poultry products; efforts to identify salmonella resistance genes in poultry could lead to more efficient poultry control strategies, thereby reducing secondary human morbidity. Genetic regulation of chicken host resistance exists, as inbred poultry lines differ in their susceptibility to infection with several strains of Salmonella. Segregation analysis with a combination of Salmonella-resistant and Salmonella-susceptible lines has shown that resistance to infection is fully dominant and is not sex-linked or associated with the major histocompatibility complex (47). The candidacy of the chicken Nramp1 homologue was tested in the differential resistance of inbred chicken lines to infection with S. Typhimurium by using sequencing and linkage analyses (48). Through the use of a mouse cDNA, the chicken homologue Nramp1 has been cloned and shown to share 68% identity with the mouse gene (49). As demonstrated in mice, the macrophage is a major site of NRAMP1 mRNA expression in chickens (49). NRAMP1 mRNA transcripts from S. Typhimurium-resistant or-susceptible chickens were analyzed to identify amino acid sequence variants that could be associated with the disease phenotype. Eleven sequence variants in Nramp1 mRNA were obtained from three Salmonella-resistant and three Salmonella-susceptible chicken lines; almost all (10) resulted in silent mutations or conservative changes (to amino acids with similar physical properties) that were detected both in resistant and susceptible chicken lines, while only one sequence variant resulted in a non-conservative substitution of a positively charged residue (Arg(sup 223) by a polar residue (Gln[sup 223] ). This allelic variant was specific to the susceptible line C and was clearly associated with survival to infection (a resistance allele at NRAMP1 improved survival rate from 13% to 27%) (48). Taken together, these data strongly suggest a direct role of NRAMP1 in susceptibility to infection in chickens. The Human NRAMP1 Gene Work in inbred strains of mice has established unambiguously that Nramp1 has an important role in determining resistance to mycobacterial infections and has encouraged several research groups to test the association of NRAMP1 with corresponding human infections. Host genetic factors play a major role in determining the outcome of mycobacterial infections in humans, as shown by racial variation in susceptibility to infection and higher concordance of tuberculosis and leprosy among monozygotic twins compared with dizygotic twins and siblings (50,51). Segregation analysis in a population from Desirade Island (French West Indies) has demonstrated that susceptibility to leprosy (regardless of the clinically defined subtype) is controlled by a major gene not linked to the major histocompatibility complex (52). Through use of a candidate gene approach, population association studies, and linkage analysis, several genes (HLA-linked genes, tumor necrosis factor, collectin, vitamin D receptor, interferon gamma receptor) have each been associated with susceptibility to mycobacterial infections (53,54). The chromosomal region surrounding Nramp1 on mouse chromosome 1 has been conserved on the telomeric end of human chromosome 2q35 and contains the human NRAMP1 orthologue (55). Sequence comparison of the mouse and human Nramp1/NRAMP1 proteins showed a high degree of conservation between the two species (85% identity, 92% similarity); the most conserved region was the intracellular loop containing the consensus sequence transport motif (56). In humans, the highest sites of NRAMP1 expression are peripheral blood leukocytes and lungs (56). The high degree of sequence homology between mouse and human NRAMP1, the presence of similar regulatory elements within the promoter regions of the genes, and similar tissue expression patterns support the notion that the NRAMP1 protein exerts similar roles in vivo in both mouse and humans. A number of polymorphic variants have been used to study the association of NRAMP1 and susceptibility to leprosy and tuberculosis (57-60). One study based on the segregation analysis of certain NRAMP1 haplotypes in 20 multiplex families involving 168 individuals from South Vietnam clearly showed that NRAMP1 was involved in predisposition to leprosy (61). Another large study measuring the association of NRAMP1 with clinical tuberculosis in a population of Gambia (West Africa) demonstrated that polymorphic variations within the human NRAMP1 gene affect susceptibility to the disease (62). Nevertheless, susceptibility to either leprosy or tuberculosis appears to be genetically heterogeneous since the role of NRAMP1 was observed only in certain ethnic groups (63,64). Identification of Nramp1 illustrates the value of comparative genomics for identification and characterization of the biologic basis for differences between susceptible and resistant hosts. Genetic dissection of the mouse model of M. bovis infection was crucial to the identification of similar mechanisms governing the human response to medically important pathogens such as tuberculosis and leprosy. Comparative genomics was also important in accelerating the identification of an important host resistance gene for salmonellosis in the chicken (a species of significant agricultural importance), where the available genetic tools are modest, relative to mice and humans. Chediak-Higashi Syndrome (CHS) CHS is a rare autosomal recessive disorder characterized by partial ocular and cutaneous albinism, a mild bleeding diathesis, and peripheral sensorimotor neuropathy. The most serious phenotype among CHS patients, however, is a marked increased in susceptibility to bacterial infection that may lead to death during the first 2 decades of life. These clinical features are attributable to dysfunctional granule-containing cells including melanocytes, platelets, Schwann cells, neurons, and granulocytes (65,66). On the basis of phenotypic similarity, the beige (bg) mutation in mice has long been regarded as a model for CHS (67). Several components of the immune system are affected in Beige/CHS. Neutrophils exhibit defective chemotaxis and reduced intracellular killing for up to 90 minutes after bacterial phagocytosis, and their granules lack the serine proteases cathepsin G and elastase because of a failure of normal protein sorting (68,69). Natural killer cell activity is defective, causing impaired cytolysis of tumors and virally infected cells; cytotoxic T-cell responses against allogeneic tumor cells are also abnormal (70,71). Mice with the bg mutation have increased susceptibility to a variety of pathogens, including cytomegalovirus, Leishmania donovani, Candida albicans, and a variety of pathogenic bacteria (E. coli, Klebsiella pneumoniae, Staphylococcus aureus, Streptococcus pneumoniae) (72-74). To identify the genetic basis of this host resistance defect, the bg gene was localized to a 0.24 cM interval of proximal mouse chromosome 13 by genetic mapping of three mouse back-crosses segregating this phenotype (75). A DNA contig of this region spanning 2,400 kb was constructed from large-capacity yeast artificial chromosomes and P1 bacteriophage clones (76). Using yeast artificial chromosome complementation and direct cDNA selection, two groups subsequently identified portions of a candidate gene for bg, named Lyst (lysosomal trafficking regulator) (77,78). Lyst, ubiquitously expressed in the mouse, has a maximum transcript size of approximately 12kb and possible complex alternative splicing. Several mutations predicted to severely truncate the Lyst polypeptide were identified within each transcript. Through the use of partial sequence data for mouse Lyst, 27 cDNAs corresponding to the human gene were identified and assembled into a complete human gene sequence of 13,499 bp, with an open reading frame of 11,403 bp (79). Comparison of the partial 3' mouse cDNA to the human sequence demonstrated 77.2% nucleotide identity and 87.9% amino acid identity, indicating that human and mouse genes are highly homologous, and sequence analysis of three CHS patients identified pathologic mutations in all. Comparative genetic mapping between the region of mouse chromosome 13 with the bg mutation and the human genome indicates homology with distal chromosome 1q. Consistent with this alignment, genetic mapping of the human CHS locus in affected families localized it to 1q42-1q44 as part of a conserved linkage group shared with mouse chromosome 13 (80,81). Radiation hybrid mapping also assigned the human CHS candidate gene to 1q43, confirming that the bg phenotype in mouse and human CHS are both caused by mutations in orthologous genes (79). Database searches with the complete nucleotide sequence of the CHS gene showed significant homology to open reading frames from S. cerevisiae and C. elegans, as well as a human cell division control protein-4 (CDC4L) (82). The modular architecture of the CHS protein is similar to Vps15, a yeast serine/threonine kinase protein kinase thought to be part of a membrane-associated signal transduction complex regulating intracellular protein trafficking (83). To date, the function of the CHS gene remains unknown, although it may be similar to Vps15 and may be part of a novel gene family. X-Linked Agammaglobulinemia (XLA) XLA, one of the first primary immunodeficiency disorders described in humans, is the prototypic example of the protective role of humoral immunity against common bacterial pathogens (84). XLA is characterized by a profound deficiency of B-lymphocyte development at two sequential stages of maturation within the bone marrow (85). This defect results in marked reductions in the serum levels of all three major classes of immunoglobulins and a profound decrease in the number of B lymphocytes in the peripheral blood as well as in the lymphoid follicles and germinal centers of lymph nodes. The clinical manifestations generally begin by the end of the first year of life, once the level of maternally derived antibodies has declined. Bacterial infections with organisms such as S. pneumoniae, Haemophilus influenzae, S. aureus, and Pseudomonas species are most common, with the respiratory tract being most frequently affected. Gastrointestinal infection with Salmonella or Campylobacter have also been reported, as have urogenital infections with Mycoplasma or Chlamydia. XLA patients have defective host resistance to enteroviruses, since neutralizing antibody is important in controlling these pathogens during their passage through the blood stream. Resistance to other infections for which intact T lymphocyte function is required (e.g., tuberculosis or histoplasmosis) remains intact. Recognition of the familial occurence of this rare disorder and pedigree analysis demonstrated an X-linked recessive inheritance pattern of the trait (86). Carrier females could not be detected because they are phenotypically normal, with normal serum levels of immunoglobulin. Linkage studies of over 500 individuals from 60 families mapped the gene for XLA to the midportion (Xq22) of the X chromosome, cosegregating with the polymorphic genetic marker DXS178 (87,88). By using complementary strategies of positional cloning and low-stringency cDNA library screening, two groups identified a novel src-like cytoplasmic tyrosine kinase, named Btk (Bruton agammaglobulinemia tyrosine kinase) as a strong candidate gene for XLA (89,90). Btk was mapped to the XLA locus by FISH and somatic cell hybrid analysis and was expressed in cell lines representing all stages of B cell development, myelomonocytic cell lines, and a macrophage cell line; it was not detectable in T lineage cell lines (90). In transformed B-cell lines from individuals affected with XLA, the expression level of Btk mRNA and protein, and consequently its kinase activity, was reduced or absent. Southern blot analysis of DNA from pedigrees with XLA cases showed restriction fragment length alterations that segregated in an X-linked recessive pattern; detailed analysis disclosed either genomic DNA deletions in the region encompassing Btk or missense point mutations resulting in nonconservative amino acid substitutions at important residues in the putative protein-tyrosine kinase domain (89). These findings provide strong evidence that the failure of normal B-cell growth and differentiation in XLA is caused by abnormal function of an intracellular protein tyrosine kinase. The CBA/N inbred mouse strain's X-linked immunodeficiency (xid) has been regarded as an experimental model for human XLA since it was first described in 1972 (91). B lymphocytes from these mice exhibit pleiotropic defects in development and function. Normal numbers of pro-B, pre-B, and surface immunoglobulin-positive B cells exist in the bone marrow, while peripheral B-cell numbers are significantly reduced (30% of normal). The B lymphocytes that are present have an abnormal surface marker phenotype, and B-cell proliferation triggered through the surface immunoglobulin M (IgM) receptor or surface immunoglobulin cross-linking is impaired, as are responses to a number of other mitogenic stimuli including lipopolysaccharide, interleukins IL-5 and IL-10, CD38 receptors, and CD40 ligands. Consistent with these defects, CBA/N mice have reduced serum IgM and IgG3 antibody levels and cannot make antibody responses when challenged with type-2 thymus-independent antigens (e.g., polysaccharides and hapten-polysaccharide conjugates). As with human XLA, impaired humoral immunity can result in increased susceptibility to bacterial pathogens, including S. Typhimurium (92). Inheritance of the susceptibility trait was linked to the xid locus by using back-cross and F2 progeny derived from crosses of CBA/N and DBA/2N parental strains. To determine whether XLA and xid were caused by mutations in homologous genes, two groups performed genetic mapping of xid and Btk. The Btk gene was closely linked to the xid locus in the distal region of the mouse X chromosome by using an interspecific back-cross mapping panel (93), and precise co-localization of Btk and xid was observed in 1,114 segregating back-cross progeny (94). Normal and mutant mouse strains did not differ in Btk expression or in vitro kinase activity. Sequence analysis of the mouse Btk transcript in CBA/N and several immunocompetent mouse strains (including the CBA/CaHN progenitor) demonstrated a point mutation within the first coding exon that is predicted to convert a highly conserved arginine residue to cysteine. This amino acid substitution occurs within the pleckstrin homology domain in the amino-terminal region of the protein and is presumed to alter normal B-cell signaling by disrupting protein-protein interactions. To unequivocally confirm that mutations in Btk were responsible for the xid phenotype, targeted gene disruption (a gene knockout experiment) was performed in embryonic stem cells (95,96). Complete elimination of Btk protein production identically reproduced the xid phenotype, indicating that the naturally occurring point mutation produces a complete loss-of-function phenotype or results in a protein with dominant negative properties (presence of a single mutant allele is sufficient to block normal gene function). The severe early B-lymphocyte developmental arrest of human XLA was not observed, which suggests that Btk function in mice is accompanied by a compensatory mechanism operating during early B-cell development to rescue B-cell maturation. On the basis of comparative mapping and sequence analysis, human XLA and the mouse xid phenotype are clearly homologous disorders caused by mutations in orthologous genes. Nevertheless, although the underlying genetic alteration in both species was successfully identified, a number of issues remain unresolved. First, the phenotypes observed in these two disorders are not identical; the more severe block of early lymphocyte development in XLA results in a greater deficiency of peripheral B cells relative to the CBA/N mouse strain, suggesting that the requirement for Btk in early murine B-cell development is less stringent than that for humans. Second, the range of pathogens to which humans are are highly susceptible appears more diverse than the range for mice. Finally, the exact role of Btk in normal B-cell physiology remains to be demonstrated. Thus far, identification of BTK has led to carrier detection and prenatal counselling; additional characterization of a mouse model with great similarity to the human condition could advance our understanding of the fundamental processes underlying B-lymphocyte development and function. Conclusions Complete understanding of infectious disease pathogenesis requires identification and characterization of host genes that regulate the response to virulent microorganisms. Through evolutionary selection, a series of innate immune defense mechanisms have evolved to protect the host against the constant threat of microbial injury and direct the development of specific adaptive immune responses. Genetic analysis of naturally occurring variation in the host response among model organisms has successfully identified novel genes such as Nramp1, Lyst, and Btk, thus providing new insights into the molecular nature of host resistance. Rapid advances are now being made in the creation and integration of dense genetic maps of model organisms and humans. Comparative genomics will play an increasingly important role in facilitating the transfer of new knowledge from experimental models to a more complete understanding of human host resistance. This work was supported by grants to Danielle Malo from the Medical Research Council of Canada (MRC), and the Canadian Bacterial Diseases Network. Dr. Qureshi is a research fellow at the Centre for the Study of Host Resistance, McGill University, and assistant physician (infectious diseases) at Montréal General Hospital. His research focuses on genetic analysis of host resistance against bacterial diseases. Address for correspondence: Danielle Malo, L11-144 Montréal General Hospital, 1650 Cedar Avenue, Montréal, Canada H3G 1A4; fax: 514-934-8261; e-mail: mc76@musica.mcgill.ca. References 1. Finlay BB, Cossart P. 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Ortega University of Arizona, Tucson, Arizona, USA --------------------------------------------------------------------------- In part, Cyclospora cayetanensis owes its recognition as an emerging pathogen to the increased use of staining methods for detecting enteric parasites such as Cryptosporidium. First reported in patients in New Guinea in 1977 but thought to be a coccidian parasite of the genus Isospora, C. cayetanensis received little attention until it was again described in 1985 in New York and Peru. In the early 1990s, human infection associated with waterborne transmission of C. cayetanensis was suspected; foodborne transmission was likewise suggested in early studies. The parasite was associated with several disease outbreaks in the United States during 1996 and 1997. This article reviews current knowledge about C. cayetanensis (including its association with waterborne and foodborne transmission), unresolved issues, and research needs. Cyclospora Overview Cyclospora cayetanensis is a protozoan parasite (subphylum Apicomplexa, subclass Coccidiasina, order Eucoccidiorida, family Eimeriidae). The organism's link to the Eimeriidae extends to the genus level through use of molecular phylogenetic analysis techniques (1). Collected data link infection to a single host—humans. In 1993, asexual meronts were described from jejunal enterocytes of humans (2). In 1997, two types of meronts and sexual stages were observed in jejunal enterocytes of biopsy specimens from infected patients excreting oocysts, confirming that the entire life cycle could be completed within a single host (3); infected persons excrete unsporulated oocysts. In the laboratory, oocysts are induced to sporulate in potassium dichromate in a petri dish at ambient temperatures (25°C to 30°C). After 1 week and up to 2 weeks, approximately 40% of oocysts contain two sporocysts with two sporozoites in each (4). Excystation of sporulated oocysts occurs in vitro when oocysts are subjected to bile salts and sodium taurocholate and mechanical pressure from a glass tube mortar and pestle (5). These findings suggest that direct person-to-person transmission is unlikely. Oocysts measure 8 µm to 10 µm in diameter and stain variably acid-fast. Without the use of an ocular micrometer, oocysts of Cyclospora might be easily confused with those of Cryptosporidium or other fecal artifacts that stain acid-fast positive, as was the case in a pseudo-outbreak of cyclosporiasis reported in Florida (6). Cyclospora oocysts are easily observed by phase contrast microscopy, and the algal-like morula appearance is evident in fresh stool specimens. A useful and distinguishing feature is oocyst autofluorescence, which appears blue by Epi-illumination and a 365-nm dichroic exciter filter and green by a 450-nm to 490-nm dichroic filter. Susceptible humans are infected by ingesting sporulated oocysts. While unknown, the infectious dose is presumed to be low. Symptoms of infection may include watery diarrhea, mild to severe nausea, anorexia, abdominal cramping, fatigue, and weight loss. Diarrhea can be intermittent and protracted (3,7,8). Persons with no previous immunity as well as very young children in developing countries are likely to exhibit symptoms. Limited data suggest that in disease-endemic countries, frequent exposure may predispose to asymptomatic infection in children and absence of infection in adults (9). Symptomatic infections can be treated with trimethoprim-sulfamethoxazole (Bactrim) (9-11). Cyclospora infections have been confirmed in North, Central, and South America, the Caribbean, England, eastern Europe, Africa, the Indian subcontinent, Southeast Asia, and Australia (12). In the United States, England, and Australia, most cases were first observed in travelers returning from the areas listed above (7,13,14). As more indigenous cases are reported from all areas, however, a cosmopolitan distribution of Cyclospora appears possible. A seasonal distribution of infection, coinciding with wet or warm months of the year, has also been suggested (15). Association with Waterborne Transmission While the organism causing Cyclospora infection was still being identified, an outbreak occurred in the staff of a Chicago hospital in 1990 (16). Infection was confirmed in 11 of 21 persons exhibiting diarrheal symptoms and lasted up to 9 weeks with alternating cycles of disease and remission. Epidemiologically, infections were associated with drinking tap water (in a resident's dormitory) possibly contaminated with stagnant water from a rooftop storage reservoir. In an isolated incident (also in Chicago), an 8-year-old child became ill and passed oocysts in the feces 1 week after swimming in Lake Michigan (7). In another isolated incident, a man from Utah became ill with severe watery diarrhea and passed oocysts after cleaning his basement, which had been flooded by sewage backup following heavy rains (8). The man's house was located near a dairy farm and much of the sewage backup was attributed to water runoff from this site. In yet another isolated incident in the United States, consumption of well water was implicated in the infection of one of three patients in Massachusetts (17). Two outbreaks of Cyclospora infection in Nepal have also been linked to waterborne transmission (18,19). In the first outbreak in 1992, expatriates, who were more likely to drink untreated water or milk reconstituted with water, became ill with diarrhea and passed oocysts. The infections occurred during the summer, which coincided with annual epidemics among the expatriates. The second waterborne disease outbreak occurred in 12 of 14 British soldiers, despite chlorination of the water involved. In this outbreak, Cyclospora oocysts were demonstrated for the first time in drinking water, which consisted of a mixture of river and municipal water. Though not directly connected with water-associated disease outbreaks, Cyclospora oocysts have been isolated from wastewater in sewage lagoons adjacent to an area of endemic disease in Lima, Peru (20); their presence was confirmed by microscopy and PCR. Water from these sewage lagoons is used to irrigate pasture land, corn fields, and trees. In other parts of Lima, water from such lagoons is used to irrigate vegetable crops. Association with Foodborne Transmission While the Nepal study conducted in 1992 strongly suggested waterborne transmission of Cyclospora, only 28% of infected patients reported drinking untreated water or milk possibly contaminated with untreated water (18). Therefore, other modes of transmission were likely, although none was identified. Foodborne transmission was suspected when consumption of raw or undercooked meat and poultry products was reported as part of case histories before the infectious organism was identified as Cyclospora (21,22). Foodborne transmission was first suggested in 1995 when the illness of an airline pilot was associated with food prepared in a Haitian kitchen and brought on board the airplane (23). Cyclospora is endemic in Haiti; this study underscored that this type of illness could be acquired from meals brought on board without visiting the country in which infection originated. Foodborne transmission of Cyclospora in the United States, first reported in 1995, was widely reported in 1996 and 1997 (24-28). Some reports early in 1996 implicated strawberries, but as more epidemiologic information was gathered, attention shifted to raspberries. In 1996, a total of 1,465 cases of cyclosporiasis were reported from 20 states (predominantly east of the Rocky Mountains), the District of Columbia, and two Canadian provinces (24). Almost half (725 cases) were event associated; the remaining (740 cases) were sporadic (i.e., not epidemiologically linked to other cases); 978 (67%) cases were laboratory confirmed; 55 clusters of cases were associated with social events. A total of 3,035 persons attended these events; 1,339 (44.1%) were interviewed, and of these 735 (54.1%) were designated case-patients. Cyclospora infection was laboratory confirmed in 238 (32.8%) cases. Raspberries were definitely served at 50 events and possibly at four more. Even in the documented 740 sporadic cases in 1996, many patients recalled eating some type of berries. Of the 54 cluster events at which raspberries were or may have been served, well-documented traceback data as to the source were uncovered for 29; of these, 21 were definitely traceable to raspberries imported from Guatemala, and an additional eight may have originated there. Twenty-five (86%) of the 29 well-documented events were traceable to one (versus more than one) exporter per event. Further tracings showed that as few as five Guatemalan farms could have accounted for the 25 events traceable to a single exporter per event. In part because of previous links with waterborne transmission, it was postulated that the berries were contaminated when sprayed with insecticides or fungicides mixed with water containing sporulated oocysts. As of August 1997, 1,450 cases of cyclosporiasis (550 laboratory confirmed) were reported (28). Many cases were cluster-associated and involved raspberries linked to Guatemala. In addition, 25 confirmed and 20 possible clusters of cases of cyclosporiasis were associated with consumption of food that contained fresh basil. An additional two clusters of cases in Florida were linked with eating mesclun lettuce (28). In each situation, the outbreaks were linked to non-Guatemalan fresh produce. Cyclospora oocysts have been isolated from vegetables from a disease-endemic area of Lima, Peru, and from Nepal (29,30). Although the number of oocysts recovered was small, encountered in only a few samples, and not associated with any known disease outbreak, the implication was clear: foodborne transmission by this route could occur. In addition, oocysts experimentally seeded on vegetables could not easily be removed by washing (30). Washing of vegetables, even though highly recommended as a means of reducing risk for infection, may therefore not totally eliminate the risk. Unresolved Issues Unresolved issues concerning Cyclospora fall into three broad categories: environmental survival, transmission to humans, and epidemiology. The boundaries of these categories frequently overlap. Environmental Survival The biggest issues of concern in this category are oocyst distribution in the environment, oocyst survival under changing conditions, and oocyst sporulation times under changing environmental conditions. All these factors affect transmission. Because of technologic limitations, Cyclospora oocysts have only been recovered in very limited numbers from water sources and vegetables (19,20,29,30). A heavy reliance has been placed on techniques used for isolating Cryptosporidium, which are inadequate (31). Very little is known about conditions that may favor the survival of Cyclospora. Preliminary studies have shown that oocysts subjected to -20°C for 24 hours and exposure to 60°C for 1 hour cannot be induced to sporulate. Oocyst storage at 4°C or 37°C for 14 days retards sporulation (32). The most intriguing environmental issue is oocyst sporulation time. The report that confirmed the identity of Cyclospora indicates that the organism requires 1 to 2 weeks to completely sporulate and become infectious under ambient conditions of 25°C to 30°C (5). Oocysts maintained at 4°C can sporulate within 6 months (4). These periods are longer than those reported for most coccidia; therefore, direct person-to-person transmission is unlikely. Also, (if confirmed under changing conditions) a prolonged sporulation time would imply that oocysts favor a moist environment, ideally water. Early in the Guatemalan berry investigations, water used to irrigate plants was thought to play a role in contaminating raspberries with oocysts. This notion, which would likely apply only to berries grown with spray irrigation, however, has largely been discarded since direct contact exposure to excessive moisture promotes rapid fruit deterioration and most raspberries grown in Guatemala rely on drip irrigation. The exact method of contamination is not known, and even though use of insecticides and fungicides made with oocyst-contaminated water has been hypothesized, its role has yet to be confirmed. If this hypothesis is true, how these agents might affect oocyst viability is also not known. Another unresolved issue is how the water might have become contaminated. Transmission to Humans The primary issues concerning transmission of Cyclospora to humans are infectious dose and species specificity. For most coccidia that infect humans and animals (e.g., Cryptosporidium [33]), the infectious dose is presumed to be low (34). What we know about the waterborne transmission of Cryptosporidium and how few of its oocysts are usually isolated from water is likely true for Cyclospora (35). However, only two foodborne outbreaks of cryptosporidiosis have been reported (one involved fresh pressed cider and the other chicken salad) (36,37). Cryptosporidium is immediately infectious upon passage from an infected person, and oocysts are usually passed in large numbers if the person is symptomatic. Unlike what has been reported for Cyclospora to date, Cryptosporidium oocysts are ubiquitous in the environment and could easily contaminate foods, especially vegetables. In one study, Cryptosporidium oocysts were recovered more frequently from vegetables than Cyclospora oocysts (30). In addition, Cryptosporidium infectious to humans has many known animal hosts (38). The issue of potential animal hosts for Cyclospora has not been resolved. Cyclospora-like organisms have been recovered from ducks, chickens, dogs, and primates (39-41). Only in primates has there been any concrete evidence identifying the agent as a species of the genus Cyclospora, and whether it is the same as C. cayetanensis is not known (41). For the other animal species mentioned, recovered oocysts, if they were oocysts of Cyclospora, may have been passing through these hosts. Attempts at finding animal hosts infected with Cyclospora-like organisms in human disease-endemic areas have largely failed, as have preliminary attempts at infecting conventionally used laboratory animals. Some researchers have convincingly shown on the basis of molecular data that Cyclospora and Eimeria are closely related (1): others have even suggested that Cyclospora should be considered a mammalian Eimeria species (42). To clarify the taxonomic issue, small subunit rRNA sequences from Isospora should be compared with those of C. cayetanensis and with Cyclospora isolates from nonhuman primates. In addition, conventional and molecular taxonomists should name the species on the basis of combined phenotypic and genotypic characteristics. Epidemiology Even though epidemiologic investigations of Cyclospora have been thorough and convincing, they raise environmental and transmission issues that require further investigation. The two areas we will consider are the relative geographic restriction of cases and attendant traceback issues associated with clusters of cyclosporiasis cases and potential indigenous infections within the United States and elsewhere. Unraveling the first issue involves tracing imported fruits or vegetables in a forward direction (possible distribution sites) as well as tracing them back (to their originating sites). In the raspberry-associated outbreaks of 1996, good traceback data were obtainable for 29 of 55 clusters. All sites (except one) were east of the Rocky Mountains. For the 25 events traceable to one (versus more than one) exporter per event, 33 (85%) of 36 shipments entered through Miami, Florida (24). If berries were also being distributed in large quantities to other, largely western regions of the country during this period, would we not expect more infections in western regions? This point, along with the attendant epidemiologic investigations, helped dissociate strawberries from reported Cyclospora infections. California strawberry growers were as likely or more likely to ship strawberries within their own region of the United States as they were to ship them elsewhere, yet most infections occurred in eastern regions of the country. In addition, Guatemalan raspberries are imported into the United States in large quantities twice a year, yet no outbreaks occurred during the winter months when this importation occurs, which indicates that the epidemiology of this infection in countries such as Guatemala where the berries are grown needs further study. The issue of indigenous U.S. infections should be investigated. Waterborne and sporadic cases have occurred in which no association could be made to raspberry consumption (7,8,16-19,24,29). Preliminary data (in one region of the United States) have linked Cyclospora infection to gardening and working with soil (43). Dr. Sterling is professor and head of the Department of Veterinary Science and Microbiology, University of Arizona, Tucson. His laboratory research focuses on cryptosporidiosis in immunologically naive and immunocompromised persons, monoclonal and polyclonal antibodies in diagnostic parasitology and immunotherapy, and Cyclospora and the microsporidia as emerging pathogens. Dr. Ortega is assistant research scientist in the Department of Veterinary Science and Microbiology, University of Arizona, Tucson. She was first to identify taxonomically Cyclospora, and her research focuses on the molecular biology and epidemiology of this organism. Address for correspondence: Charles R. Sterling, Department of Veterinary Science and Microbiology, University of Arizona, Tucson, AZ 85721, USA; fax: 520-621-2799; e-mail: csterlin@u.arizona.edu. References 1. Relman DA, Schmidt TM, Gajadhar A, Sogin M, Cross J, Yoder K, et al. Molecular phylogenetic analysis of Cyclospora, the human intestinal pathogen, suggests that it is closely related to Eimeria species. J Infect Dis 1996;173:440-5. 2. Bendall RP, Lucas S, Moody A, Tovey G, Chiodini PL. Diarrhoea associated with cyanobacterium-like bodies: a new coccidian enteritis of man. Lancet 1993;341:590-2. 3. Ortega YR, Nagle R, Gilman RH, Watanabe J, Miyagui J, Quispe H, et al. Pathologic and clinical findings in patients with cyclosporiasis and a description of intracellular parasite life-cycle stages. J Infect Dis 1997;176:1584-9. 4. Ortega YR, Sterling CR, Gilman RH, Cama VA, Diaz F. Cyclospora species—a new protozoan pathogen of humans. N Engl J Med 1993;328:1308-12. 5. Ortega YR, Sterling CR, Gilman RH. A new coccidian parasite (Apicomplexa:Eimeriidae) from humans. J Parasitol 1994;80:625-9. 6. Sterling CR, Ortega YR, Hartwig EC, Pawlowicz MB, Cook MT, Miller JR, et al. Outbreaks of pseudo-infection with Cyclospora and Cryptosporidium—Florida and New York City, 1995. MMWR Morb Mortal Wkly Rep 1997;46:354-8. 7. Wurtz R. Cyclospora: a newly identified intestinal pathogen of humans. Clin Infec Dis 1994;18:620-3. 8. Hale D, Aldeen W, Carroll K. Diarrhea associated with Cyanobacteria-like bodies in an immunocompetent host. An unusual epidemiological source. JAMA 1994;271:144-5. 9. Madico G, Gilman RH, Cabrera L, Sterling CR. Epidemiology and treatment of Cyclospora cayetanensis infection in Peruvian children. Clin Infect Dis 1997;24:977-81. 10. Pape JW, Verdier RI, Boncy M, Boncy J, Johnson W. Cyclospora infection in adults infected with HIV. Clinical manifestations, treatment, and prophylaxis. Ann Intern Med 1994;121:654-7. 11. Hoge CW, Shlim DR, Ghimire M, Rabold JG, Pandey P, Walch A, et al. Placebo-controlled trial of co-trimoxazole for Cyclospora infections among travelers and foreign residents in Nepal. Lancet 1995;345:691-3. 12. Soave R. Cyclospora: an overview. Clin Infect Dis 1996;23:429-37. 13. Bendall RP, Chiodini PL. The epidemiology of human Cyclospora infection in the UK. In: Betts WB, Casemore D, Fricker C, Smith H, Watkins J, editors. Protozoan parasites and water. Cambridge: The Royal Society of Chemistry, Thomas Graham House; 1995. p. 26-9. 14. McDougall TJ, Tandy MW. Coccidian/cyanobacterium-like bodies as a cause of diarrhea in Australia. Pathology 1993;25:375-8. 15. Hoge CW, Echeverria P, Rajah R, Jacobs J, Malthouse S, Chapman E, et al. Prevalence of Cyclospora species and other enteric pathogens among children less than 5 years of age in Nepal. J Clin Microbiol 1995;33:3058-60. 16. Huang P, Weber JT, Sosin DM, Griffin PM, Long EG, Murphy JJ, et al. The first reported outbreak of diarrheal illness associated with Cyclospora in the United States. Ann Intern Med 1995;123:409-14. 17. Oii WW, Zimmerman SK, Needham CA. Cyclospora species as a gastrointestinal pathogen in immunocompetent hosts. J Clin Microbiol 1995;33:1267-9. 18. Hoge CW, Shlim D, Rajah R, Triplett J, Shear M, Rabold JG, et al. Epidemiology of diarrhoeal illness associated with coccidian-like organism among travelers and foreign residents in Nepal. Lancet 1993;341:1175-9. 19. Rabold JG, Hoge CW, Shlim DR, Kefford C, Rajah R, Echeverria P. Cyclospora outbreak associated with chlorinated drinking water [letter]. Lancet 1994;344:1360-1. 20. Sturbaum GD, Ortega YR, Gilman RH, Sterling CR, Klein DA. Detection of Cyclospora cayetanensis in sewage water. Appl Environ Microbiol 1998;64:2284-6. 21. Ashford RW. Occurrence of an undescribed coccidian in man in Papua New Guinea. Ann Trop Med Parasitol 1979;73:497-500. 22. Hart AS, Ridinger MT, Soundarajan R, Peters CS, Swiatlo AL, Kocka E. Novel organisms associated with chronic diarrhea in AIDS. Lancet 1990;335:169-70. 23. Connor BA, Shlim DR. Foodborne transmission of Cyclospora. Lancet 1995;346:1634. 24. Herwaldt BL, Ackers M-L, and the Cyclospora working group. An outbreak in 1996 of cyclosporiasis associated with imported raspberries. N Engl J Med 1997;336:1548-58. 25. Jacquette G, Guido F, Jacobs J, Smith P, Adler D. Update. Outbreaks of cyclosporiasis—United States, 1997. MMWR Morb Mortal Wkly Rep 1997;46:461-2. 26. Hofman J, Liu Z, Genese C, Wolf G, Manley W, Pilot K, et al. Update: outbreaks of Cyclospora cayetanensis infection—United States and Canada. MMWR Morb Mortal Wkly Rep 1996;45:611-2. 27. Chambers J, Somerfeldt S, Mackey L, Nichols S, Ball R, Roberts D, et al. Outbreaks of Cyclospora cayetanensis infection—United States, 1996. MMWR Morb Mortal Wkly Rep 1996;45:549-51. 28. Pritchett R, Gossman C, Radke V, Moore J, Busenlehner, Fischer K, et al. Outbreak of cyclosporiasis. Northern Virginia-Washington, DC.-Baltimore, Maryland, Metropolitan Area, 1997. MMWR Morb Mortal Wkly Rep 1997;46:689-91. 29. Kocka F, Peters C, Dacumos E, Azarcon E, Kallick C, Langkop C. Outbreaks of diarrheal illness associated with cyanobacteria (Blue-green algae)-like bodies-Chicago and Nepal, 1989 and 1990. MMWR Morb Mortal Wkly Rep 1991;40:325-7. 30. Ortega YR, Roxas CR, Gilman RH, Miller NJ, Cabrera L, Taquiri C, et al. Isolation of Cryptosporidium parvum and Cyclospora cayetanensis from vegetables collected from markets of an endemic region in Peru. Am J Trop Med Hyg 1997;57:683-6. 31. Steiner TS, Thielman NM, Guerrant RL. Protozoal agents: what are the dangers for the public water supply? [review] Annu Rev Med 1997;48:329-40. 32. Smith HV, Paton CA, Mtambo MMA, Girdwood RWA. Sporulation of Cyclospora sp oocysts. Appl Envir Microbiol 1997;63:1631-2. 33. DuPont H, Chappell CL, Sterling CR, Okhuysen PC, Rose JB, Jakubowski W. The infectivity of Cryptosporidium parvum in healthy volunteers. N Engl J Med 1995;332:855-9. 34. Jackson GJ, Leclerc JE, Bier JW, Madden JM. Cyclospora—still another new foodborne pathogen. Food Technology 1997;51:120. 35. Rose JR, Lisle JT, LeChevallier M. Waterborne cryptosporidiosis: incidence, outbreaks, and treatment strategies. In: Fayer R, editor. Cryptosporidium and cryptosporidiosis. Boca Raton (FL): CRC Press, Inc.; 1997. p. 93-109. 36. Besser-Wiek JW, Forfang J, Hedberg CW, Korlath JA, Osterholm MT, Sterling CR, et al. Foodborne outbreak of diarrheal illness associated with Cryptosporidium parvum—Minnesota, 1995. MMWR Morb Mortal Wkly Rep 1996;45:783-4. 37. Millard PS, Gensheimer KF, Addiss DG, Sosin DM, Houck-Jankoski A, Hudson A. An outbreak of cryptosporidiosis from fresh pressed apple cider. JAMA 1994;272:1592-6. 38. Meng J, Doyle MP. Emerging issues in microbiological food safety [review]. Annu Rev Nutr 1997;17:255-75. 39. Garcia-Lopez HL, Rodriguez-Tovar LE, Medina-de la Garza CE. Identification of Cyclospora in poultry [letter]. Emerg Infect Dis 1996;2:356-7. 40. Yai LE, Bauab AR, Hirschfeld MP, de Oliveira ML, Damaceno JT. The first two cases of Cyclospora in dogs, Sao Paulo, Brasil. Rev Inst Med Trop Sao Paulo 1997;39:177-9. 41. Smith HV, Paton C, Girdwood RAW, Mtambo MMA. Cyclospora in non-human primastes in Gombe, Tanzania. Vet Rec 1996;138:528. 42. Pieniazek NJ, Herwaldt BL. Reevaluating the molecular taxonomy: is the human associated Cyclospora a mammalian Eimeria species? Emerg Infect Dis 1997;3:381-3. 43. Koumans EH, Katz D, Malecki J, Wahlquist S, Kumar S, Hightower A, et al. Novel parasite and mode of transmission: Cyclospora infection—Florida. 1996. Annual Epidemic Intelligence Service Conference 1996;45:60. —————————————————————————————————————————————————————————————————————————— Synopses Using Monoclonal Antibodies to Prevent Mucosal Transmission of Epidemic Infectious Diseases Larry Zeitlin,* Richard A. Cone,*† and Kevin J. Whaley *† *ReProtect, LLC, Baltimore, Maryland, USA; and †The Johns Hopkins University, Baltimore, Maryland, USA --------------------------------------------------------------------------- Passive immunization with antibodies has been shown to prevent a wide variety of diseases. Recent advances in monoclonal antibody technology are enabling the development of new methods for passive immunization of mucosal surfaces. Human monoclonal antibodies, produced rapidly, inexpensively, and in large quantities, may help prevent respiratory, diarrheal, and sexually transmitted diseases on a public health scale. In 1975, Köhler and Milstein noted that monoclonal antibodies (MAbs) "...could be valuable for medical and industrial use" (1). Since then, the use of MAbs has become routine in the research and diagnostic laboratory, but antibodies have yet to be used to their maximum potential in medical and public health applications. Two recent reviews of the therapeutic use of antibodies suggest that systemically administered antibodies may play an important role in treating infections by drug-resistant pathogens as well as pathogens for which no antimicrobial drugs are available (2,3). However, the greatest potential for MAbs probably lies in prevention since antibodies are in general more effective for prophylaxis than for therapy (3,4). From a public health perspective, prevention is especially important (5). In particular, direct application of MAbs to mucosal surfaces blocks the entry of pathogens into the body. We review here the evidence of antibody efficacy in preventing disease and recent advances that have facilitated the development of MAbs for mucosal applications in humans. Finally, we consider the public health potential of topical delivery of MAbs for preventing mucosal transmission of infections. Immunologic Strategies for Preventing Mucosal Transmission Vaccines that stimulate systemic immunity can prevent systemic disease, but generally fail to prevent mucosal disease. Vaccines that stimulate active mucosal immunity have demonstrated good efficacy in animal models, but with few exceptions (polio and influenza vaccines), have not been as effective as they could be in humans. Some of the discrepancies between study results in animals and humans are probably due to a failure of studies in animals to model immune evasion strategies of pathogens (6) that occur in humans. These strategies include rapid evolution of variable strains (7), pathogens that coat themselves with host antigens (8), and pathogens that are transmitted to a new host by hiding inside cells shed by the infected host (cell vectors) (9). Furthermore, most vaccines successful in stimulating mucosal immunity in animals contain irritating adjuvants or attenuated pathogens, which are generally considered unacceptable for use in humans; vaccines with human-safe adjuvants have not generated high concentrations of protective antibody in the mucosa. Current research is investigating improved immunogens, delivery vehicles, and adjuvants, as well as exploring the best inductive sites for generating a protective mucosal immune response at a specific mucosal surface (10). In contrast to vaccines, passive immunizations can deliver protective levels of antibodies immediately and directly to the susceptible mucosal surface (Figure 1-top). Also, with passive mucosal immunization, it may be possible to defeat some key immune evasion strategies by using antibodies directed against host cell vectors, host antigens that coat the pathogen, or receptors used by pathogens to enter target cells (11). In addition, new methods for the sustained release of antibodies offer the possibility of long-term protection (12). [Fig] Figure 1. Topical delivery of pathogen-specific MAbs can protect the mucosal epithelium. (Top) Protective MAbs (in this figure, secretory immunoglobulin A; SIgA) can be topically applied to the mucosa in various ways. (Bottom) In mucus, MAbs are believed to act by a number of mechanisms to prevent penetration of the mucous layer and subsequent infection of target cells (62). MAbs can trap pathogens in the mucous gel by forming low affinity bonds with mucin fibers and can agglutinate pathogens into clusters too large to diffuse through the mucous gel. Efficacy of Antibodies in Preventing Disease The first use of immune serum for preventing disease by passive immunization was reported more than 100 years ago by von Behring and Kitasato (13). Subsequently, systemic passive immunization with antibodies has been proven effective in preventing many diseases. By binding to a pathogen, systemically delivered antibodies can inhibit attachment to and fusion with target cells, inhibit internalization by target cells, inhibit uncoating inside a cell, aggregate pathogens thereby preventing them from reaching target cells, interact with complement to lyse the pathogen, induce phagocytosis of the pathogen, and cause killer cells to lyse the pathogen by antibody-dependent cellular cytotoxicity (14). Table 1 lists the highest efficacy reported for systemically delivered antibodies in preventing disease in mammalian species and against a wide range of pathogens that infect humans. No antiviral treatments are available for most viruses listed in the table, yet antibodies can prevent the diseases caused by all of these viruses. Table 1: Examples of highly effective systemic passive immunization ----------------------------------------------------------------------- Anti- Species body Preven- Pathogen (sup a) (sup b) tion(%) DRS(sup c) Ref. ----------------------------------------------------------------------- Viruses Chikungunya mou p 100 (15) Cytomegalovirus hum p 50 X (16) Dengue mou p 100 (17) Ebola bab p 80 (18) Hantavirus mou m 100 (19) Herpes simplex (genital) mou m 100 X (20) (ocular) mou m 100 (21) HIV mou m 100 X (22) Hepatitis A hum p 90 (23) Hepatitis B hum p 92 (24) Influenza mou m 100 (25) Lassa mon p 100 (26) Measles mou m 100 (27) Polio hum p 58 (28) Rabies mou m 100 (29) antibodies Reovirus mou m 100 (30) Rift Valley fever ham p 100 (31) Respiratory syncytial hum m 100 (32) p 40 (33) Rubella hum p 57 (34) Varicella zoster hum p 100 (35) Venezuelan equine mou p 100 (36) encephalomyelitis Bacteria Borrelia burgdorferi ham p 100 (37) Bordetella pertussis mou m 100 X (38) Chlamydia pneumoniae mou p 100 (39) Chl. trachomatis mou m 90 (40) Escherichia coli rat m 100 X (41) Francisella tularensis mou p 100 (42) Group B Streptococcus mou m 100 X (43) Haemophilus influenzae rat p 100 X (44) Mycoplasma pneumoniae ham p 80 (45) Neisseria meningitis mou m 90 X (46) Proteus mirabilis mou m 100 X (47) Pseudomonas aeruginosa mou p 100 X (48) Salmonella Typhimurium mou p 100 X (49) Shigella flexneri rab p 100 X (50) Staphylococcus aureus rab m 100 X (51) Streptococcus pneumoniae mou p 90 X (52) Treponema pallidum ham p 100 (53) Yersinia pestis mou p 100 (54) m NR(sup d) (55) Fungi Candida albicans mou p > 67 X (56) Cryptococcus neoformans mou m 70 X (57) Parasites Plasmodium falciparum mon p 75 X (58) Toxoplasma gondii mou m 100 (59) ----------------------------------------------------------------------- (sup a)Species: mou=mouse; hum=human; bab=baboon; mon=monkey; ham=hamster; rat= rat; rab=rabbit. (sup b)Antibody: m=monoclonal; p=polyclonal. (sup c)DRS=Drug-resistant strains reported (from Ref. 60). (sup d)NR = not reported Although less studied than systemic passive immunization, the prophylactic use of mucosal antibodies predates the therapeutic use of immune sera. Antibodies delivered in mother's milk have been protecting the gastrointestinal tract of nursing infants since the mammary gland first evolved approximately 50 million years ago. Most infections begin in mucosal surfaces (approximately 400 m(sup 2)in an adult human); supplementing the antibody repertoire in a mucous secretion (Figure 1-top) thus offers an effective method for protecting a mucosal surface against pathogens to which the host has not been exposed or become immune. In addition to the protective mechanisms described above, antibodies delivered to mucosal surfaces can trap pathogens in the mucous gel, make them mucophilic, and prevent their diffusion and motility (Figure 1-bottom); as a result, pathogens trapped in mucus are shed from the body with the normal flow of mucous secretions or are digested if these secretions enter the digestive tract (61-63). Topical passive immunization of mucosa can block transmission of bacteria, viruses, fungi, and parasites that infect humans (Table 2). Table 2: Examples of highly effective topical passive immunization of mucosa herpes simplex ----------------------------------------------------------- Anti- Pre- Species Route body ven- Pathogen (sup a) (sup b)(sup c)tion Ref. ----------------------------------------------------------- Viruses Herpes simplex mou v m 100% (64,65) r m 100% (66) Influenza fer o p 100% (67) mou n p >4(sup d) (68) Rotavirus hum o p 100% (69,70) Respiratory syncytial mon n m 3-4(sup d) (71) Bacteria Chlamydia trachomatis mou v m 90% (72) Clostridium difficule ham o p 100% (73) Escherichia coli hum o p 100% (74) Porphyromonas gingivalis hum o m 100% (75) Shigella flexneri hum o p 100% (76) Staphylococcus aureus mou n p 3-4(sup e) (77) Streptococcus mutans hum o m 100% (78) Vibrio cholerae mou o m 100% (79) Fungi Candida albicans mou v p >50(sup f)(80) Parasites Cryptosporidium parvum mou o m 77(sup g)(81) ----------------------------------------------------------- (sup a)Species tested in: mou=mouse; fer=ferret; hum=human; mon=monkey; ham=hamster. (sup b)Delivery route of pathogen and antibody: v=vaginal; r=rectal; o=oral; n=nasal. (sup c)Antibody: m=monoclonal; p=polyclonal. (sup d) log10 reduction in virus titer. (sup e) log10 reduction in cfu. (sup f) % reduction in cfu. (sup g) % reduction in number of parasites. The predominant (and perhaps the most appropriate for mucosal delivery) antibody isotype on most human mucosal surfaces is secretory immunoglobulin A (SIgA); efficient methods for producing SIgA have been reported (82,83). SIgA, a tetravalent dimer of monomeric IgA associated with two polypeptides (joining chain and secretory component), is especially stable and well suited to function in the enzymatically hostile environment that prevails at mucosal surfaces (84). SIgA, the least phlogistic class of antibody (84), is the least likely to induce inflammatory responses that can make it easier for toxins and pathogens to breach the mucosal surface. Immune exclusion of antigens, enzymes, and toxins has been repeatedly demonstrated in vivo, and protection generally correlates with levels of SIgA antibodies in the relevant mucous secretions. Finally, the protective role of SIgA has been demonstrated in many systems (85). Recent Advances in mAb Technology Generating High-Affinity Human MAbs Since the advent of cloning of human antibodies from combinatorial libraries constructed from seropositive persons (86,87), generation of fully human MAbs against human pathogens has become routine (Figure 2) (88). For example, from a single bone marrow donor, human MAbs were prepared against HIV, respiratory syncytial virus cytomegalovirus, herpes simplex virus types 1 and 2, varicella zoster virus, and rubella virus (88). MAbs can even be obtained from naive libraries prepared from unexposed persons (if the large enough repertoire) (89); therefore, antibodies against pathogens lethal to humans can be generated. Alternatively, human MAbs can be generated by traditional immunization of commercially available mice that have been genetically engineered to contain human immunoglobulin loci in their germline (Figure 2)(90,91). [Fig] Figure 2. Generation of human monoclonal antibodies. (Phage display) Heavy and light chain cDNA isolated from human B-cells is used to generate a combinatorial library in which random heavy (H) and light chain (L) pairings are expressed on the surface of phage. These phage can then be screened for antigen binding by traditional techniques (e.g., ELISA). Since only the antigen binding region is used in the phage display process, the selected clone is then placed into an appropriate expression vector to produce a full antibody molecule. (Transgenics) Genetically manipulated mice have been produced with inactivated endogenous immunoglobulin genes, and with unrearranged human immunoglobulin gene segments introduced (90,91). These mice are then immunized with antigen, and hybridomas are produced by traditional routes. (See refs. 88, 89 for more technical information on these two methods and refs. 92, 93 for comparisons of these two methods). Dramatic enhancement of the affinity of an mAb has been demonstrated by molecular biologic techniques in which mutants of an antibody are generated and then screened for higher affinity or higher neutralization activity (93-95). For example, the affinity of one anti-HIV mAb has been enhanced 420-fold, and this matured antibody neutralizes more HIV strains than the original mAb (94). Furthermore, expressing a mAb as a multivalent isotype, such as SIgA or IgM, can dramatically enhance the potency of an antibody by increasing the avidity (96) or agglutination activity (14). For example, an anti-Escherichia coli IgM was 1,000-fold more effective in protecting neonatal rats than its class-switched IgG (both in vitro and in vivo)(41). From a commercial standpoint, a 1,000-fold increase in avidity could translate into a 1,000-fold decrease in dose and subsequent cost. Also, a large dose of a highly potent mAb can substantially increase the duration of protection (97). Production Systems MAbs have traditionally been produced in cell culture and have been prohibitively expensive for most preventive uses. Over the years, however, the cost has continually dropped; MAbs are now being produced in cell culture for $200 to $1,000 per gram (98,99). Production of MAbs has recently been reported in both transgenic plants and animals (82,100,101). Both of these systems are expected to lower costs dramatically. Indeed, transgenic plants can be scaled up in agricultural fields to produce tons of "plantibody," and plant-produced antibody is predicted to cost less than U.S. $1/g (102). The actual cost, however, will remain unknown until large-scale batches are produced, purified, and formulated in accordance with Good Manufacturing Practices. Safety and Regulatory Status More than 80 MAbs are now in clinical trials (most for cancer imaging or therapy) and more than one quarter of these are in phase III trials (103). Few safety problems have been reported for systemic applications; antibodies are now considered "biotechnology-derived pharmaceuticals" by the U.S. Food and Drug Administration (FDA)—enabling a more straightforward regulatory process than in the past (92,104). Even though MAbs have often been evaluated for systemic applications, only recently have they been evaluated in humans for mucosal applications. This new interest in mucosal antibodies may be partially due to the increasing recognition of the importance of mucosal immunity. Only two clinical trials have evaluated topically delivered MAbs: intranasally delivered anti-RSV in infants at high risk (105) and orally delivered anti-Streptococcus mutans in adults (106); no major adverse effects were reported in these studies. Safety concerns, such as peptide and glycosylation immunogenicity, are important when MAbs are delivered systemically but are likely to be of less concern when MAbs are applied to the mucosa, a surface that has evolved to interact with the external environment. Indeed, antibodies delivered to the lumen of a mucosal surface have minimal interaction with circulating immune cells. Although proteins, and even antibodies, can be absorbed through mucosal surfaces (107,108), generally only small quantities are absorbed (109,110). The inability of SIgA to activate complement by the classic pathway is likely involved in maintaining the integrity of mucosal surfaces (63); therefore, SIgA may be preferable to IgG or IgM for many mucosal applications. The FDA "Points to Consider" for characterization of antibodies produced in cell-culture and transgenic animals (111) are better defined than for characterization of antibodies produced in transgenic plants; however, plant-derived antibodies are free of animal viruses and may therefore not require rigorous viral inactivation processing steps. In addition, although glycosylation patterns of MAbs produced in mammalian cell-culture and transgenic animals are closer phylogenetically to humans than glycosylation patterns in plants, given our repeated exposure to plant sugars in food and personal care products, it is unlikely that any of these patterns are novel to human immune systems (112). In fact, in a recently completed clinical trial with repeated applications of plant-produced antibody for the prevention of oral colonization by S. mutans, no safety problems were encountered, nor were there any detectable human anti-plant antibody responses (113). Selection for resistant organisms by widespread and repeated use of antibiotics is a serious health concern (60). Drug-resistant strains of a wide variety of pathogens have already been reported (Table 1). Antibiotic or antiviral treatment of infected persons in which pathogens are actively replicating provides a strong evolutionary selection process for developing drug-resistant pathogens. In contrast, MAbs are less likely to create resistant organisms when used in a preventive context at a mucosal surface against a pathogen that is not yet actively replicating. Even if a systemic infection does occur during topical use of MAbs, resistant organisms will likely not be created since the pathogen will not be replicating and evolving in the presence of the mAb applied to the mucosal surface. This is in marked contrast to the settings in which antibiotics and antiviral drugs select for resistant strains (60). If MAbs are used frequently on a population level, the risk of selecting for resistant organisms may increase. When the emergence of resistant strains is of particular concern, the tendency to select mAb-resistant organisms could be minimized by using cocktails of mucosal antibodies directed at multiple antigenic targets (2,114). Because new MAbs can be produced with a rapid turnaround time (discussed below), the emergence of an antibody-resistant strain could be countered by producing a new mAb directed toward the mutated epitope or another antigenic target of the resistant strain. Indeed, the flexibility of the antibody structure to create a virtually inexhaustible repertoire of antigen binding specificities suggests that immunoglobulins evolved in part as a means to cope rapidly with new pathogens. Turnaround Time for Developing a New mAb Since human MAbs can be identified quickly by cloning variable regions from specific antigen-binding human lymphocytes (115) or panning combinatorial libraries (87), antibodies could be used as a rapidly developed method for defending against new pathogens. The time required for collecting lymphocytes from a seropositive person, screening for an appropriate antibody, cloning, and expressing the antibody in culture in a well-equipped laboratory is 1 to 3 months; quantities sufficient for protecting persons at high risk or those at the focal point of an outbreak could be available in fewer than 6 months. High-capacity production in quantities sufficient for broad public health application could be available in several years, assuming that the safety of antibodies as a class of molecules is established and an infrastructure is in place for producing these antibodies. While in rare instances vaccines can be developed this quickly (e.g., the 1976 influenza vaccine [5]), new vaccines, antibiotics, and antiviral therapies usually take considerably longer to develop. Moreover, even though passive immunization may require repeated applications, MAbs delivered to a mucosal surface can provide immediate protection against infection. Potential Preventive Uses for Topically Delivered MAbs From a public health perspective, MAbs are most promising for preventing gastrointestinal, respiratory, and reproductive tract infections. These infections cause almost 11 million deaths annually worldwide, accounting for more than 50% of the deaths caused by communicable diseases and 22% of deaths by all causes (116). Sexually transmitted diseases (STDs) accounted for 87% of all cases reported among the top ten most frequently reported diseases in 1995 in the United States; more than 12 million Americans are infected with STDs each year at an estimated annual cost of more than $12 billion (117). If a track record of safety and efficacy can be achieved, mucosal antibodies will probably be most useful as over-the-counter products that could reach populations not well integrated into the health-care system. The condom, a nonmedical over-the-counter personal protection product, has played an important preventive role in the HIV epidemic. Personal protection provided by over-the-counter antibody-based technology could play a similar role in future emerging disease epidemics. Diarrheal Disease Studies in animal models have demonstrated that orally delivered antibodies were 100% effective in preventing rotavirus (70) and cholera (79) infections. In humans, orally delivered bovine antibodies were 100% effective in preventing rotavirus (118), enterogenic E. coli (74), Shigella infection (76), and necrotizing enterocolitis (119). For orally delivered MAbs, digestive degradation is a potential concern. However, significant levels of functional antibody survive treatment with pepsin at pH 2 or with a pool of pancreatic enzymes at pH 7.5 in vitro (120). In addition, most ingested IgA in milk survives passage through the gastrointestinal tract of infants (121); intact antibody delivered orally with an antacid survived passage through the gastrointestinal tract of adults (74,76). Assuming that a 10-mg dose of antibody is protective (i.e., assuming that the mAb is only 100-fold more potent than polyclonal preparations [118]), the production costs for the amount of plantibody needed for 100 days of protection could be approximately one cent (102). Since diarrheal diseases are most prevalent in developing countries, preventive strategies must be extremely inexpensive; therefore, MAbs produced in plants or in the milk of animals are likely most suitable for these countries. Because of the speed with which MAbs pass through the gastrointestinal tract, antibodies delivered orally will need to be delivered frequently, perhaps more than once a day. In endemic-disease regions, MAbs could be delivered orally as a supplement with food or water. Respiratory Disease Animal studies have demonstrated the efficacy of nasal delivery of antibodies for the prevention of RSV infection (71) and influenza (68). In one study, topical application was approximately 100 times more effective than systemic delivery (122). Another study found an anti-RSV mAb (MEDI-493) to be approximately 100 times more effective than an equal quantity of a polyclonal preparation (32). These results suggest that 10,000 times less anti-RSV mAb would be required for topical applications than for systemically delivered polyclonal preparations. Protective systemic doses of MEDI-493 are approximately 100 mg (15 mg/kg) (32), so <1 mg might suffice for protection if this mAb were applied topically. Intranasally applied mAb has a residence half-time of a little under one day in the monkey (71), suggesting that once-a-day applications that deliver several-fold more than a protective dose can provide continuous protection. MAbs for protecting the respiratory tract could be delivered in nose drops or by aerosol once a day to those at particular risk (e.g., infants and the elderly during influenza season) or to everyone living near the epicenter of an epidemic. STDs With the exception of hepatitis B, no vaccines are available for the prevention of STDs (Table 3). Until effective and safe vaccines are developed, vaginal delivery of a cocktail of anti-STD pathogen MAbs might make an effective new method for broad spectrum protection against STDs (11). In animal models, MAbs have been shown to protect against transmission of C. albicans, C. trachomatis, HSV, HIV, and syphilis (Tables 1, 2) (11). Antibodies have been delivered experimentally to the vagina in solution, gels, and more recently, by sustained release devices for long-term delivery of protective MAbs (123,124). Antibodies were found to be stable when stored in seminal fluid or cervical mucus for 48 hours at 37°C (125); no significant inactivation occurred over the pH range of the human vagina (pH 4 to 7) for at least 24 hours at 37°C (Zeitlin et al., unpub. obs.). Since the effective half-life of antibodies applied topically depends on the turnover time of mucus, a single vaginal application may thus provide protection for at least 1 day, and probably several days (97). If so, passive immunization of the vagina may extend protection to the occasional days when the user forgets to apply the mAb. Considering there are an estimated 5 billion acts of sexual intercourse per year in the United States (11), large-scale production of MAbs in plants may offer the best system for the low costs needed for such a public health initiative. In addition, because the most common class of infection in the first month of life is primarily caused by STD pathogens present in the birth canal (126), the same mucosal antibodies could be used in a predelivery cervicovaginal lavage or applied to newborns' eyes for studies in the prevention of ophthalmia neonatorum. Indeed, in some cultures the mother's colostrum, a fluid rich in SIgA, is applied to the newborns' eyes (127). Table 3: Preventive vaccines or cures for major sexually transmitted disease pathogens ---------------------------------------------------------- Pathogen Vaccine Cure DRS(sup a) ---------------------------------------------------------- Chlamydia trachomatis no yes Haemophilus ducreyi no yes X Hepatitis B yes no Herpes simplex 1 and 2 no no X HIV-1 and 2 no no X Human papilloma virus (HPV) no yes(sup b) Neisseria gonorrhoeae no yes X Treponema pallidum no yes Trichomonas vaginalis no yes X ---------------------------------------------------------- (sup a)Drug-resistant strains reported. (sup b)Surgical removal of HPV-infected tissue is performed. HPV-related cervical cancer identified early has a high cure rate; however, in the United States, for every three new cases, there is approximately one death (117). Conclusions In animal models and human studies, antibodies have been shown to prevent a wide variety of infectious human diseases. Recent advances allow development of a new era of mucosal mAb-based products. These advances include the development of combinatorial libraries for rapid selection of human MAbs, the ability to increase dramatically the potency of a specific mAb, and the marked reduction in the cost of cell-culture—produced MAbs as well as the ability to produce MAbs inexpensively and at high capacity in transgenic animals and plants. In addition, since MAbs can be developed considerably more rapidly than most vaccines and antimicrobial drugs, MAbs may prove useful for combating emerging pathogens. Mucosal infections account for a large percentage of infectious disease-related illness and deaths; hence topical passive immunization with MAbs may offer a new opportunity for improving public health. Finally, many of the remaining safety issues regarding the human use of mucosal MAbs are likely to be addressed by clinical trials now under way. Acknowledgments The authors thank the Rockefeller Foundation Bellagio Study and Conference Center and Drs. Polly F. Harrison, Mich B. Hein, and Thomas R. Moench for their review of drafts. Dr. Zeitlin is a research scientist at ReProtect, LLC. His interests focus on the development of monoclonal antibodies for contraception and the prevention of sexually transmitted diseases. Address for correspondence: Kevin Whaley, 3400 North Charles St. Jenkins Hall, Baltimore, MD 21218, USA; fax: 410-516-6597; e-mail: whale@jhu.edu. References 1. Kohler G, Milstein C. Continuous cultures of fused cells secreting antibody of predefined specificity. Nature 1975;256:495-7. 2. Casadevall A, Scharff M. Return to the past: the case for antibody-based therapies in infectious diseases. Clin Infect Dis 1995;21:150-61. 3. Casadevall A. Antibody-based therapies for emerging infectious diseases. Emerg Infect Dis 1996;2:200-8. 4. Cross A. Intravenous immunoglobulins to prevent and treat infectious diseases. 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Bandea,* James Baggs,‡ and Danuta Pieniazek* *Centers for Disease Control and Prevention, Atlanta, Georgia, USA; †Universidade Federal do Rio de Janeiro, Brazil; ‡Emory University, Atlanta, Georgia, USA --------------------------------------------------------------------------- We systematically evaluated multiple and recombinant infections in an HIV-infected population selected for vaccine trials. Seventy-nine HIV-1 infected persons in a clinical cohort study in Rio de Janeiro, Brazil, were evaluated for 1 year. A combination of molecular screening assays and DNA sequencing showed 3 dual infections (3.8%), 6 recombinant infections (7.6%), and 70 (88.6%) infections involving single viral subtypes. In the three dual infections, we identified HIV-1 subtypes F and B, F and D, and B and D; in contrast, the single and recombinant infections involved only HIV-1 subtypes B and F. The recombinants had five distinct B/F mosaic patterns: B(sup gag-p17)/B(sup gag -p24)/F(sup pol(sup)/B(sup env) , F(sup gag-p17)/B(sup gag -p24)/F(sup pol)/F(sup env) , B(sup gag-p17)/B-F(sup gag -p24)/F(sup pol)/F(sup env) , B(sup gag-p17)/B-F(sup gag -p24)/F(sup pol)/B(sup env) , and F(sup gag-p17)/B-F(sup gag -p24)/F(sup pol)/F(sup env). No association was found between dual or recombinant infections and demographic or clinical variables. These findings indicate that dual and recombinant infections are emerging as an integral part of thE HIV/AIDS epidemic in Brazil and emphasize the heterogenous character of epidemics emerging in countries where multiple viral subtypes coexist. Our understanding of the global molecular epidemiology of HIV-1 infections has improved substantially in recent years. Remarkable antigenic diversity, especially in the viral envelope, has emerged among HIV isolates worldwide (1). As the HIV/AIDS pandemic grows, viral strains are becoming more geographically dispersed, and the simultaneous presence of multiple subtypes in a given region is now common (2). Mixed infections and recombinants involving sequences of distinct HIV-1 subtypes (mosaics) are being recognized, but their prevalence and effect on the pandemic have not been fully evaluated (3). Consequently, the distribution of dual infections and mosaic viruses among different populations and the changes of this distribution over time are still relatively unknown. In addition, scientists are increasingly interested in possible differences in the transmission, epidemiologic patterns, and natural history of HIV-1 infections caused by more than one viral subtype and recombinant genomes. Finally, the efficacy of HIV-1 vaccines, primarily developed against subtype B viruses, may differ against more divergent recombinant variants and mixed infections of distinct HIV-1 subtypes. Such knowledge is vital to understanding the relevant role of mixed infections as a prerequisite for recombination and could be applied immediately in molecular epidemiology and immunotherapy. Studies using convenience samples first documented mixed infections caused by viruses of subtypes B and E in Thailand (4). Subsequently, such studies have identified cases of dual infections with subtypes B, F, C, and D in Brazil (5-7), B and F in Puerto Rico (8), A and C in Rwanda (9), and triple HIV-1 infection with groups O and M of different clades in a single Cameroonian AIDS patient (10). In addition, potential dual infections have been detected by molecular screening assays among HIV-1 infected populations in Uganda and Kenya, where subtypes A and D coexist (11). The consequences of HIV-1 mixed infections may profoundly influence the dynamic of the pandemic through altered patterns of viral transmission and pathogenesis. Moreover, the resulting genetic variation may lead to the emergence of new HIV variants, including those with altered antigenicity and reduced sensitivity to detection by current diagnostic assays. The global emergence of such variants is exemplified in the impact of two HIV-1 recombinants of subtypes A/E and G/A on epidemics in Thailand and in certain parts of Central Africa, respectively (1,12-14). Interestingly, the presumptive parental subtypes E and G have not been identified. Because Brazil has been selected as a World Health Organization field site for HIV-1 vaccine evaluation programs, priority has been given to extensive molecular examination of the prevalence and genetic diversity of HIV-1 strains circulating in the country. By November 1997, 116,277 AIDS cases had been reported to the Brazilian AIDS Control Program of the Ministry of Health (15). The 293 HIV-1 strains that have been molecularly characterized document that, although four HIV-1 subtypes——B, F, C, and D are——circulating in Brazil, only subtypes B and F are common (16-21). Moreover, five HIV-1 dual infections were identified in 21 HIV-infected patients during testing of molecular techniques that would discriminate between HIV-1 infections caused by single and multiple subtypes (5-7). Also, two cases of B/F recombinants have been found accidentally through sequence analysis of the env region (22). These data only indicate the potential for dual and recombinant infections in Brazil, where multiple subtypes circulate; they cannot, however, be used to assess the frequency of these infections among the HIV-infected population. In this study, we evaluated the proportion of HIV-1 dual and recombinant infections among 79 patients enrolled in a prospective clinical cohort study in Rio de Janeiro (an area where HIV-1 subtypes B and F are common). The Study Population Part of an ongoing prospective clinical cohort study established in 1991 by the AIDS program of the Federal University of Rio de Janeiro, the HIV-1-infected patients have been continuously enrolled in the cohort for consultation, treatment, evaluation of different clinical parameters during the progression of the disease, and assessment of the genetic variation of HIV-1 strains (23). With informed consent, blood samples were obtained from the 79 patients, who consecutively attended the clinic between October and December 1994. For the purposes of this study, samples collected in 1994 were compared as needed, and follow-up specimens were collected 1 year later. The clinical profile of patients was based on the medical evaluation at the time of blood collection. Because the patients were randomly selected, the findings presented in this article likely reflect the trends in the HIV/AIDS epidemic in the Rio de Janeiro area. Epi Info, Version 6 program (CDC, Atlanta, GA, USA) was used to calculate frequencies, means, and analyses of variance of demographic information, clinical stages, and laboratory parameters. Design To investigate the proportion of mixed infections, we must distinguish between those involving two or more distinct HIV-1 strains and infection with a single intersubtype recombinant strain. By definition, the mixed infection occurs when multiple phylogenetically distinct copies of the same gene representing different viral genomes are present within one patient (24). In contrast, in the mosaic strain, different viral regions within the same genome are classified by phylogenetic analysis into different subtypes (3). Potential HIV-1 mixed infections involving distinct HIV-1 variants were segregated from single infections caused by only one subtype by using a restriction fragment length polymorphism (RFLP) screening assay of the prt gene (5,7). The restriction map profiles of the viral prt allow the segregation of HIV-1 strains to subtypes A, B, C, D, and F (Figure 1A). AluI digestion patterns separate subtypes A, C, and F from subtypes B and D. Sequential restriction analysis of prt using HinfI, BclI, and ScaI restriction enzymes further differentiates among these two subtype groups. The simultaneous occurrence of more than one digestion pattern indicates a potential mixed infection (Figure 1B). We selected recombinants among RFLP-subtyped single infections by additionally subtyping the C2-V3 env region with the heteroduplex mobility assay (25). Subtype discrepancies between prt and env regions were considered potential HIV-1 recombinants. All potential multiple infections and recombinants were analyzed by sequence analysis. The search for recombinants was further expanded by sequence analysis of the entire p17 gag or a 311-bp of the p24 gag fragment or both among preselected prt-env recombinants, all prt-env subtype F, and some prt-env subtype B viruses. [Fig] Figure 1. Differentiation between single (A) and dual (B) HIV-1 infections by the restriction fragment length polymorphism analysis of the polymerase chain reaction–amplified prt. A: Three AluI digestion patterns represent subtypes A, C, and F (pattern 1) and subtypes B and D (patterns 2 and 3); two HinfI patterns represent subtypes D (pattern 1) and B (pattern 2); two BclI patterns represent subtypes F (patterns 1) and A and C (pattern 2); two ScaI patterns represent subtypes A (pattern 1) and C (pattern 2). B: Two AluI digestion patterns (1 and 2) in the dually infected patient with HIV-1 subtypes F and B; two HinfI patterns (1 and 2) in the patient infected with subtypes B and D viruses. Polymerase Chain Reaction (PCR) Uncultured or cultured peripheral blood mononuclear cells from patients were used for nested-PCR of the entire HIV-1 protease gene (prt, 297-bp), p24 gag fragment (311-bp), and C2-V3 domain of env (565-bp) (6,7). The outer primers for amplification of a 717-bp gag fragment spanning the entire 17 gag region and a 311-bp of p24 gag were LTRF: 5'GGGCTAATTTGGTCAAAAAGAAG; nucleotide position: 6-28, HIV-1(sub MN), and P24RA: 5'ATGTCACTTCCCCTTGGTTCT; nucleotide position: 1482-1502. The inner primers were P17F: 5'GCAAGAGGCGAGGGGCAGCAGCCG; nucleotide position: 716-739, and P17R: 5'CCCATTCTGCAGCTTCATTGA; nucleotide position: 1413-1433. The outer primers for amplification of a 1,444-bp fragment from p24 gag to the reverse transcriptase region were P24F1: 5'ATAGAGGAAGAGCAAAACAAAA; nucleotide position: 1,099 to 1,120, and MOPR1: 5'AAAATTGGAGTATTGTATGGATT; nucleotide position: 2,724 to 2,746. The inner primers were P24FA: 5'CAAAATTACCCTATAGTGCA; nucleotide position: 1,177 to 1,196, and MOPR2: 5'GGTCCATCCATTCCTGGTTT; nucleotide position: 2,601 to 2,620. PCR conditions were the same for amplification of all viral regions (7). Sensitivity of Detection of Dual Infections To determine the sensitivity of detection of dual infections by the RFLP assay, we mixed 5, 10, 25, 50, and 100 copies of subtypes B and F cloned proviral DNA in equal proportions or in ratios 1:2, 1:4, 1:10, and 1:20 for nested PCR amplification of prt (Table 1). For each combination, one to four independent PCR reactions were run. The simultaneous amplification of the viral prt of subtypes B and F was evaluated by the RFLP assay. PCR controls included DNA templates of single HIV-1 subtypes. Table 1. Sensitivity of detection of HIV-1 dual infections caused by viruses of subtypes and F ----------------------------------------------------------------- No. of viral subtypes No. of F B F:B experiments B F B&F ----------------------------------------------------------------- 100 100 1:1 1 – – + 50 50 1:1 4 – – + 25 25 1:1 4 – + (1) +(3) 10 10 1:1 4 – – + 5 5 1:1 4 – + (3) + (1) 100 5 20:1 2 – + (1) + (1) 100 10 10:1 2 – + (1) + (1) 100 25 4:1 2 – + – 100 50 2:1 2 – – + 5 100 1:20 1 – – + 10 100 1:10 1 – – + 25 100 1:4 1 – – + 50 100 1:2 1 – – + 100 0 3 – + – 50 0 3 – + – 5 0 3 – + – 10 0 3 – + – 5 0 2 – + – 0 100 3 + – – 0 50 2 + – – 0 25 3 + – – 0 10 3 + – – 0 5 3 + – – ------------------------------------------------------------------ Absence (-) and presence (+) of AluI digestion pattern of prt characteristic for subtype B, subtype F, and combination of subtypes B and F (Fig. 1). The cloned proviral DNA of HIV-1 subtypes B and F spanning a 1444-bp fragment from p24 gag to rt was used for the nested PCR amplification of prt. Amplified products were digested with AluI restriction enzyme, and the presence of two digestion patterns was analyzed on a 10% polyacrylamide gel by ethidium-bromide staining. Cloning, Sequencing, and Phylogenetic Analysis The PCR-amplified proviral prt sequences from potential dual infections were cloned by using the Original TA Cloning Kit (Invitrogen, Carlsbad, CA, USA). DNA from 30 clones of each specimen Was screened for distinct HIV-1 sequences by the RFLP assay (7). Double-stranded viral DNA from selected clones or from direct PCR-amplified prt, p17 and p24 gag, and C2-V3 env products was cycle-sequenced in both directions with fluorescent dye–labeled sequencing terminators (26). Sequencing reactions were run in an automated DNA sequencer (Applied Biosystems, Foster City, CA, USA). The sequences were aligned by the CLUSTAL multiple sequence alignment program (27). After gaps were eliminated, the aligned sequences were analyzed by the maximum likelihood method, with the fastDNAml program, which uses randomized data input and global rearrangement (28). Additionally, the neighbor joining method (PHYLIP package version 3.5c [29]) was used, with or without bootstrapping. The stability of the tree's topology was tested by pruning (removing one sequence from the alignment and rerunning the phylogenetic analysis). The SIV-cpz sequences (GenBank accession no. X52154) were used as outgroups. HIV-1 sequences generated in this study have been submitted to GenBank. Results Detection of Mixed Infections RFLP analysis of the prt gene showed the simultaneous presence of two different digestion patterns in specimens from 10 of 79 patients. A complex pattern composed of elements of AluI patterns #1 (subtypes A, C, or F) and #2 (subtypes B or D) was identified in nine patients, and a combination of two HinfI digestion patterns (subtypes B and D) was found in one patient (Figure 1B). These data suggest that each of 10 patients could be infected with multiple distinct HIV-1 subtypes. The remaining 69 samples were classified as single infections caused by viruses of prt subtypes B (59) and F (10). The PCR-amplified viral prt products from these 10 patients were cloned and sequenced to confirm the presence of mixed infections. Sequence analysis showed the simultaneous presence of two distinct HIV-1 variants in only three patients: BR45, BR62, and BR83. The nucleotide divergence between the two prt sequences within the patients was 6.8% for BR45, 7.4% for BR62, and 7.1% for BR83, indicating two distinct HIV-1 variants in each of these patients (6). Phylogenetic analysis confirmed these findings and demonstrated that the divergent HIV-1 prt sequences segregated into subtypes B and F (BR45), subtypes D and F (BR62), and subtypes B and D (BR83) (Figure 2a). In the remaining seven specimens, the observed RFLP results were consequences of either point mutations in AluI restriction site (five cases) or G A hypermutation (two cases), which occurred across one of the sequences within each specimen and destroyed the defining AluI sites. These changes in AluI restriction sites gave rise to genetically distinct quasispecies within the patients but did not represent distinct subtypes, as further confirmed by phylogenetic analysis (e.g., BR55-1 and BR55-2, and BR99-1 and BR99-2 in Figure 2a). Thus, despite the presence of mixed AluI digestion patterns in these seven specimens, they were classified as single infections of subtype B variants. [Fig] Figure 2. Phylogenetic classification of HIV-1 sequences from Brazilian patients (denoted with BR prefix). The trees were constructed on the basis of DNA sequences of prt (a), env (b), gag-p24 (c), and gag-p17 (d) by the neighbor-joining method. Numbers at the branch nodes connected with subtypes indicate bootstrap values. An arrow indicates dually infected specimens; an asterisk shows viral sequences, which clustered into different lineages depending on which parts of viral genome were analyzed; ! represents hypermutated sequences; ? represents unclassified subtype of p24 gag sequences. The distinct HIV-1 subtypes are delineated. The scale bar indicates an evolutionary distance of 0.10 nucleotides per position in the sequence. Vertical distances are for clarity only. GenBank accession numbers: prt [AF099155-99171;AF079986-79989;AF079991; and AF079994-79996]; env [AF113560-113576]; p17gag [AF115443-115451]; p24gag [AF115780-115797]. To address the issue of potential laboratory contamination, we collected repeat blood samples from dually infected patients approximately 1 year later and processed them on separate occasions. (Blood was unlikely to be contaminated during collection because a disposable vacutainer system was used to obtain each blood sample.) The sequence data from the first and second blood samples of each person showed a 98% to 99% similarity. Also, the viral sequences from these patients were distinct from those of laboratory strains (Figure 2a; 855M, 8,986, and 9,001) commonly used as standards. Sensitivity of Detection of Dual Infections To investigate the sensitivity of the RFLP screening method, we performed reconstruction experiments, in which two distinct viral DNA templates of subtypes B and F were analyzed in the same reaction mixture (Table 1). When equal proportions of 10 to 100 HIV-1 DNA template copies were used for prt amplification, two viral subtypes could be simultaneously detected in all but one of 19 experiments. However, when five or fewer copies of each viral subtype were used, two subtypes were identified in only one of four assays; in the remaining three assays either subtype B or subtype F amplicons, but not both, were found. Similarly, in experiments containing varying proportions of subtypes B and F DNA templates, the simultaneous presence of two viral subtypes could be detected in only 8 (66%) of 12 experiments. In comparison, 5 to 100 copies of a single viral subtype (B or F) were routinely amplified and identified in all control reactions. Detection of Intersubtype HIV-1 Recombinants Parallel RFLP/heteroduplex mobility assay screening for HIV-1 subtypes in the prt and env regions identified two potential recombinants among 76 single infections. Subtype F prt and subtype B env were found in both specimens BR43 and BR60 during this initial screening. The remaining specimens were classified into subtypes F (n = 8) and B (n = 66) in both prt and env regions. These prt-env potential recombinants, all subtype F specimens, and eight selected subtype B samples were further evaluated by sequence analysis, which confirmed the results of the screening assays (Figure 2a, 2b). The search for the HIV-1 recombinant genome in these 18 samples was further expanded to the gag region. Mosaic sequences of subtypes B and F were found within the p17 or p24 gag (Table 2, Figure 2, discussion below) in four prt-env subtype F variants (BR46, BR57, BR59, and BR97). Similarly, in one of two prt-env recombinants (BR60), the gag sequences had a mosaic pattern. In contrast, the p24 gag sequences of eight prt-env subtype B specimens were homogeneous and also classified as subtype B (Figure 2). Taken together, the comparative molecular analysis of gag, pol, and env regions allowed the identification of six specimens that carried HIV-1 recombinant genomes, representing five distinct mosaic structures: B(sup gag-p17)/B(sup gag -p24)/F(sup pol)/B(sup env) , F(sup gag-p17)/B(sup gag -p24)/F(sup pol)/F(sup env) , B(sup gag-p17)/B-F(sup gag -p24)/F(sup pol)/F(sup env) , B(sup gag-p17)/ B-F(sup gag -p24)/F(sup pol)/B(sup env) , and F(sup gag-p17)/B-F(sup gag -p24)/F(sup pol)/F(sup env) . Table 2. p17/p24 gag, prt, and C2-V3 genetic subtyping of HIV-1 DNA sequences(sup a) from peripheral blood mononuclear cells collected from 18 patients in Rio de Janeiro ------------------------------------------------------------------- Genotypes Specimen No. gag p17 gag p24 prt C2-V3 ------------------------------------------------------------------ Group 1(sup b) 8 ND(sup c) B B B Group 2(sup d) 4 F F F F (BR46) (NA[sup e]) (B) (F) (F) (BR59) (F) (B) (F) (F) (BR57) (F) (B/F) (F) (F) (BR97) (B) (B/F) (F) (F) (BR60) (B) (B/F) (F) (B) (BR43) (B) (B) (F) (B) ------------------------------------------------------------------ (sup a)Recombinant specimens are shown in parentheses; B/F mosaic structure within a 311-bp of the p24 gag fragment consisting of subtypes B and F sequences. (sup b)Group 1: BR34, BR52, BR55, BR64, BR65,BR71, BR75, and BR92. (sup c)ND=not done. (sup d)Group 2: BR41, BR54, BR58, and BR112. (sup e)NA = not available due to negative PCR. [Fig] Figure 3. Analysis of the putative recombination within the gag region. The aligned sequences were classified into subtype B, subtype F, and recombinant subtype B/F on the basis of linearity of subtype assignment for the p17-p24 gag region. Asterisks show characteristic nucleotide patterns for subtypes B and F sequences; dots represent nucleotides homologous to the MN gag sequence; dashes indicate gaps introduced to maintain the alignment; and arrows indicate the potential recombination regions within the p24 gag fragment. The nucleotide position is marked. The potential crossover breakpoints within 717-bp of the p17-p24 gag mosaic sequences were examined by comparison with nucleotide signature patterns characteristic for subtypes B and F viruses (Figure 3). We performed comparative analyses with aligned DNA sequences of recombinants (BR57, BR59, BR60, and BR97), subtype B (MN and BR43), and subtype F variants (BR41, BR54, BR58, and BR112). This analysis confirmed an intragene recombination within p24 gag in specimens BR57, BR60, and BR97—a finding consistent with our failure to phylogenetically assign these gag sequences to any known subtype (Figure 2c). The putative breakpoints within the intragene recombinant sequences were located between nucleotides 97 and 137 in sample BR57 and between nucleotides 173 and 213 in BR60 and BR97 (Figure 3). The exact breakpoint position could not be determined because of extensive sequence homology between subtypes B and F in this viral region. This analysis also revealed putative crossover breakpoints for variants BR57 and BR59 in proximity to the coding region for the p17-p24 protein-processing site. The potential breakpoints between the second half of gag and the beginning of pol region in variants BR43, BR46, and BR59 are being investigated. To examine the possibility of in vitro recombination during PCR amplification (30), we performed PCR amplification of the long fragments covering the gag and prt area in the endpoint- diluted lysates (31). To ensure that endpoint PCR products were amplified from single copy templates, we used samples only from dilutions at which 1 of 10 PCR amplifications were productive for further sequence analysis. The comparative analysis of the entire p17 gag, p24 gag, and prt sequences demonstrated 98% homology between the undiluted and diluted lysates—strong evidence that the recombinant sequences were not a result of the PCR amplification process. Epidemiologic and Clinical Characteristics The mean ages of patients infected with HIV-1 of single subtype B or F were not different from those with dual or recombinant infections (p = 0.77) (Table 3). In addition, the patients did not differ significantly by gender, risk group, and clinical stage of disease (p = 0.44 to p = 0.48). The patients infected with HIV-1 subtype B (403) and subtype F (854) did, however, differ (p = 0.04) by mean CD4 counts. Although dates of seroconversion were not known for all patients, the earliest HIV-positive results were reported among patients infected with subtype B (in agreement with previous observations that the spread of subtype B viruses occurred earlier than other HIV-1 subtypes in Brazil [16,17]), which might explain the difference in CD4 counts. Table 3. Characteristics of the study population -------------------------------------------------------------------------- Dual Recombinant infection infection Subtype B Subtype F Characteristic n=3 n=6 n=66 n=4 -------------------------------------------------------------------------- Gender (Female/male) 1/2 3/3 24/42 3/1 Mean age in years 36 (30-44) 34 (25-45) 37 (23-60) 34 (26-43) (range) Clinical stage(sup a) 1 3 5 29 4 2 – 1 17 – 3 – – 14 – 4 – – 6 – Mean CD4 cells 527 484 403 823 (range) (404-743) (190-821) (59-1281)(sup b) (570-1270) Years of 1st serologic tests 1985-1987 – – 2 – 1988-1991 2 2 30 2 1992-1994 1 4 34 2 Heterosexual 2 5 27 4 Homosexual 1 – 19 – Bisexual – – 8 – Blood transfusion – – 3 – recipient Intravenous drug user – – 1 – Multiple factors – 1 2 – Unknown – – 6 – -------------------------------------------------------------------------- (sup a)WHO staging system [32]. (sup b)Based on data available for 54 patients. Conclusions HIV-1 infections caused by dual and intersubtype-recombinant genome may be relatively common among HIV-1-infected Brazilians. Using both heteroduplex mobility assay and RFLP screening methods, as well as sequencing, we identified three dual (3.8%) and six recombinant (7.6%) infections involving distinct viral subtypes among 79 HIV-1-infected persons from Rio de Janeiro. We chose viral prt to screen for dual infections because this highly conserved region provides the best opportunity for simultaneous amplification of distinct HIV-1 variants in the same PCR reaction. Proviral prt sequences can be routinely amplified from approximately 95% of all analyzed seropositive samples collected from the Americas, Asia, Africa, and Europe (data not shown). Moreover, the RFLP assay of the prt gene is convenient for screening a large number of samples (5). The detection of B/F, B/D, and F/D dual infections is in agreement with our 1993 study among patients from the same Rio de Janeiro cohort, which showed five dual HIV-1 infections involving subtypes F and B (one case), F and D (one case), and B and C (three cases of familial clustering) among 21 HIV-infected persons (5-7). Interestingly, dual infections involving HIV-1 subtype D continue to be detected in patients from Rio de Janeiro, an area with a high predominance of HIV-1 subtypes B and F, but rare subtype D, single infections. Because retrospective specimens (peripheral blood mononuclear cells) were not available for these patients, we could not confirm whether the acquisition of mixed strains was sequential (superinfection) or simultaneous. However, combination of subtypes F or B with rare subtype D viruses among some dual infections may suggest that in these cases two viral strains might be acquired through cotransmission rather than through superinfection. All naturally occurring HIV-1 dual infections are likely not detected by current methods. First, quantitative differences in two distinct viral DNA templates in the sample can lead to selective PCR amplification of only one subtype. Second, despite targeting conserved genes such as prt, some divergent viral strains may escape PCR amplification because of primer mismatches. Finally, a single nucleotide mutation in the endonuclease restriction site can abrogate the recognition pattern and distort detection of dual infections in the RFLP analysis. These observations suggest that the rate of HIV-1 mixed infections within this Brazilian cohort might be even higher than 4%. Our previous findings of five dual infections among 21 patients from the same cohort support this assumption. If we take into account these five cases, the percentage of mixed infections caused by viruses of distinct subtypes circulating between 1993 and 1994 among 100 patients analyzed from this Rio de Janeiro cohort would increase to 8%. The potential underestimate of mixed infections is highlighted by the additional detection of 6 (7.6%) distinct recombinants within the cohort. This finding is consistent with the estimated 5% to 10% intersubtype mosaics among HIV-1 genomes in the Los Alamos database (3). Interestingly, all recombinant or mosaic genomes described in this report involved only subtypes B and F viral regions, although dual infections caused by other subtypes were also circulating in this cohort. Moreover, our results indicate that recombination between gag and pol (prt) regions is more frequent than between pol and env and lead to speculation that such stable B/F mosaics have selective advantage. Such observations support the assumption that recombination within the gag gene occurs more often than within other viral regions (33) and emphasize the need for rapid subtyping methods specific to the gag region. To investigate the potential impact of dual and recombinant infections on the clinical status of patients, we compared the clinical and demographic characteristics of these patients with those of patients infected with one nonrecombinant viral subtype. Epidemiologic information for all 79 patients was available only at the first draw of blood. Although the results did not show significant differences between the two groups, the possibility of differences exists. Our findings provide the first baseline measure of the range of HIV variability in Rio de Janeiro from 1993 to 1994 and the proportion of dual and recombinant infections among the HIV-infected Brazilian population. Future systematic molecular epidemiologic surveys of HIV heterogeneity in Rio de Janeiro may show the potential changes in the molecular profile of the HIV/AIDS epidemic over time; the laboratory tools we used to identify single, dual, and recombinant infections may be useful in such investigations. Nevertheless, our study on genetic variation of HIV-1 subtypes among blood donors from the state of Rio de Janeiro documented the presence of mosaic viruses of subtypes B and F and subtypes B and D in blood units collected in 1996 (34). Moreover, recent genetic analysis of viral strains collected in 1997 from HIV-1-infected patients living in Manaus (a city in Brazil's Amazon region) showed the presence of dual infections and recombinants caused by subtypes B and F viruses (A. Tanuri, pers. comm.). Therefore, these data indicate that the heterogenic pattern of HIV-1 infections, first observed in Rio de Janeiro, also exists in other regions of Brazil. 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Cornelissen M, Kampinga G, Zorgdrager F, Goudsmit J, and the UNAIDS Network for HIV Isolation and Characterization. Human immunodeficiency virus type 1 subtypes defined by env show high frequency of recombinant gag genes. J Virol 1996;70:8209-12. 34. Tanuri A, Swanson P, Devare S, Berro OJ, Savedra A, Costa LJ, et al. HIV-1 subtypes among blood donors from Rio de Janeiro, Brazil. J Acquired Immune Defic Syndr. In press 1998. —————————————————————————————————————————————————————————————————————————— Research Genetic Diversity and Distribution of Peromyscus-Borne Hantaviruses in North America Martha C. Monroe,* Sergey P. Morzunov,† Angela M. Johnson,* Michael D. Bowen,* Harvey Artsob,‡ Terry Yates,§ C.J. Peters,* Pierre E. Rollin,* Thomas G. Ksiazek,* and Stuart T. Nichol* *Centers for Disease Control and Prevention, Atlanta, Georgia, USA; †University of Nevada, Reno, Nevada, USA; ‡Laboratory Centre for Disease Control, Federal Laboratories, Winnipeg, Manitoba, Canada; and §University of New Mexico, Albuquerque, New Mexico, USA --------------------------------------------------------------------------- The 1993 outbreak of hantavirus pulmonary syndrome (HPS) in the southwestern United States was associated with Sin Nombre virus, a rodent-borne hantavirus; The virus' primary reservoir is the deer mouse (Peromyscus maniculatus). Hantavirus-infected rodents were identified in various regions of North America. An extensive nucleotide sequence database of an 139 bp fragment amplified from virus M genomic segments was generated. Phylogenetic analysis confirmed that SNV-like hantaviruses are widely distributed in Peromyscus species rodents throughout North America. Classic SNV is the major cause of HPS in North America, but other Peromyscine-borne hantaviruses, e.g., New York and Monongahela viruses, are also associated with HPS cases. Although genetically diverse, SNV-like viruses have slowly coevolved with their rodent hosts. We show that the genetic relationships of hantaviruses in the Americas are complex, most likely as a result of the rapid radiation and speciation of New World sigmodontine rodents and occasional virus-host switching events. Hantaviruses, rodent-borne RNA viruses, can be found worldwide. The Old World hantaviruses, such as Hantaan, Seoul, and Puumala, long known to be associated with human disease, cause hemorrhagic fever with renal syndrome of varying degrees of severity (1). After hantavirus pulmonary syndrome (HPS) was discovered in the southwestern United States in 1993 (2-4), intensive efforts were begun to detect and characterize hantaviruses in North America and determine their public health importance (5). As of January 1999, 205 HPS cases had been confirmed in 30 states of the United States, and 30 cases had been confirmed in three provinces of Canada; most cases occurred in the western regions of both countries. While Sin Nombre virus (SNV) has been identified as the cause of most HPS cases in North America, an increasingly complex array of additional hantaviruses has appeared (Table 1). Table 1. Hantaviruses in the New World --------------------------------------------------------------------------- Virus(sup a)Disease(sup b)Known or suspected host Location Virus isolate --------------------------------------------------------------------------- Sigmodontinae associated West & Sin Nombre HPS Peromyscus maniculatus Central Y (grassland form) U.S.and Canada Eastern U.S. Monongahela HPS P. maniculatus (forest form) and Canada N New P. leucopus (eastern York HPS haplotype) Eastern U.S. Y Blue River P. leucopus (SW/NW haplotypes) Central U.S. N Bayou HPS Oryzomys palustris Southwestern Y U.S. Black Creek Sigmodon hispidus (eastern Canal HPS form) Florida Y Muleshoe S. hispidus (western form) Southern N U.S. Caño Delgadito S. alstoni Venezuela Y Andes HPS Oligoryzomys longicaudatus Argentina Y and Chile Oran HPS O. longicaudatus Northwestern N Argentina Central Lechiguanas HPS O. flavescens Argentina N Bermejo O. chacoensis Northwestern N Argentina Hu39694 HPS Unknown Central ArgentinaN Pergamino Akadon azarae Central N Argentina Maciel Bolomys obscurus Central N Argentina Laguna Paraguay and Negra HPS Calomys laucha Bolivia Y Juquitiba HPS Unknown Brazil N Rio Mamore O. microtis Bolivia and Y Peru El Moro Western U.S. Canyon Reithrodontomys megalotis and Mexico N Rio Segundo R. mexicanus Costa Rica N Arvicolinae associated Prospect Hill Microtus pennsylvanicus N. America Y Bloodland M. ochrogaster N. America N Lake Prospect M. Hill-like pennsyl./montanus/ochrogaster N. America N Isla Vista M. californicus Western U.S. N and Mexico Murinae associated Seoul HFRS Rattus norvegicus Worldwide Y --------------------------------------------------------------------------- (sup a)Major virus types or species are in bold and indented below the rodent subfamilies with which they are associated; related genetically distinct virus lineages that may represent additional species or subspecies are indented below virus types and species. (sup b)HPS = hantavirus pulmonary syndrome; HFRS = hemorrhagic fever with renal syndrome. Surveys of rodents for hantavirus antibody have shown hantavirus-infected rodents in most areas of North America (3;6-9; Ksiazek et al., unpub. data; Artsob et al., unpub data). Serologic evidence of hantavirus infection has been found in North American rodents of the family Muridae. Most North American hantaviruses are associated with the subfamily Sigmodontinae; only a small number are associated with the subfamilies Arvicolinae or Murinae. To determine the number and distribution of hantaviruses in North America, we conducted a nucleotide sequence analysis of a polymerase chain reaction (PCR) fragment amplified from a large number of representative HPS patient and hantavirus-infected rodent samples from throughout the region. We focused on the North American viruses (particularly those associated with Peromyscus species rodents), although the nucleotide sequences of many hantaviruses from South America and elsewhere were included as outgroups to increase the resolution of the analysis. Genetic Detection and Phylogenetic Analysis of New World Hantaviruses The nucleotide sequences of 139 bp fragments of the G2 encoding region of virus M segments amplified by reverse transcriptase-PCR (RT-PCR) from 288 hantavirus-infected rodent and human samples were compiled from Genbank sources or from data reported here. Details of the specimen selection and methods of genomic analysis are provided in the Appendix. The Genbank accession numbers of those sequences published earlier (bigtree.xls) can be accessed here. The entire aligned dataset (bigtree.nex), including 130 newly presented sample sequences, is also available here. These sequences include those derived from 229 SNV-like viruses associated with Peromyscus species rodents from throughout North America. Maximum parsimony analysis of the aligned sequences was conducted with PAUP (12; Appendix), which resulted in a reasonably well-defined tree topology with several distinct lineages of SNV-like viruses and other clearly discernable hantaviruses (Figures 1, 2). Bootstrap analysis showed that while several of the major nodes of the tree were not well supported (values of 50% or less), many others were robust (values of 70% or higher) (Figures 1, 2). In most phylogenetic analyses, bootstrap values provide highly conservative estimates of the probability of correctly inferring the corresponding clades (13). Bootstrap values of 70% or higher corresponded to a probability of 95% or higher that the corresponding clade was correctly identified. Values of 50% or lower corresponded to a probability of 65% or lower that the clade was correctly identified (13). [fig 1] Figure 1. Overall hantavirus phylogenetic tree based on analysis of a 139 nucleotide fragment of the G2 coding region of the virus M segment. All newly reported sequences are shown bolded. The three virus groups corresponding to the hantaviruses carried by rodents of the subfamilies Murinae, Sigmodontinae and Arvicolinae are indicated. P indicates the clade containing the Sin Nombre- like viruses found in Peromyscus species rodents, the details of which are shown in figure 2. Horizontal branch lengths are proportional to the nucleotide step differences between taxa and predicted nodes. No scale bar is indicated as the actual number values are arbitary due to the weighting used in the successive approximations method (see appendix for details). Bootstrap values greater than 50% are indicated above branches. Virus labels include the virus or virus lineage name (ISLA, Isla Vista; TULA, Tula; PH, Prospect Hill or Prospect Hill-like; KBR, Khabarovsk; PUU, Puumala; SN, Sin Nombre; ELMC, El Moro Canyon; CDG, Caño Delgadito; BCC, Black Creek Canal; BAY, Bayou; JUQ, Juquitiba; AND, Andes; LN, Laguna Negra; THAI, Thailand; DOB, Dobrava; HTN, Hantaan; SEO, Seoul), species source of material (Mcalif, Microtus californicus; Mross, Microtus rossiaemeridionalis; Marv, Microtus arvalis; Mpenn, Microtus pennsylvanicus; Mmont, Microtus montanus, Mochro, Microtus ochrogaster; Mfort,Microtus fortis; Cg, Clethrionomys glareolus; Rm, Reithrodontomys megalotis;Salst, Sigmodon alstoni; Shisp, Sigmodon hispidus; Op, Oryzomys palustris; Cal, Calomys laucha; Bi, Bandicota indica; Apfl, Apodemus flavicollis; Apag, Apodemus agrarius; Rn Rattus norvegicus), identifier, and state, region or country of origin. For spreadsheet containing the details of all samples, click on (bigtree.xls) [fig 2] Figure 2. Phylogenetic tree of hantaviruses associated with Peromyscus species rodents. Figure provides a detailed view of clade P in figure 1. S indicates clade containing classical SN virus samples detected in humans or P. maniculatus. See figure 1 legend for overall tree description. Additional species source of material abbreviations include: Pm, Peromyscus maniculatus; Pl, Peromyscus leucopus, Prg.fasc, Perognathus fasciatus; Tam.quad, Tamias quadrimaculatus; Pt, Peromyscus truei; Mus musc., Mus musculus, and Tam.dors., Tamias dorsalis. Samples from historic materials are followed by an H. Diversity of New World Hantaviruses As expected on the basis of earlier nucleotide sequence analysis of a limited number of complete S or M hantavirus genome segments or virus genome fragments (5), the evolutionary relationships among hantaviruses were closely correlated with those of their known or suspected primary rodent reservoirs (Figure 1; Table 1). Hantaviruses associated with subfamily Murinae rodents (Hantaan, Dobrava, Seoul, and Thailand viruses) are clearly separated from those associated with Arvicolinae and Sigmodontinae rodents. The Arvicolinae-associated viruses (Puumala, Khabarovsk, Tula, Isla Vista, Prospect Hill [PH], and PH-like viruses]) form a reasonably well-supported clade, but the phylogenetic position of this group relative to the Murinae- and Sigmodontinae-associated viruses is not well resolved. The New World hantaviruses of the Arvicolinae group, primarily associated with Microtus species voles, include not only the classic PH virus (labeled PHV-1), originally isolated from M. pennsylvanicus in Maryland (14,15), and two other distinct PH-like virus lineages recently found in this vole species in North Dakota (R737 and R731; R742), but also Isla Vista virus in M. californicus, PH-like hantavirus lineages in M. ochrogaster in North Dakota (R812 and R789), and M. montanus in Wyoming and Nevada (3485; LY-R2312) (16,17). Virus phylogenetic placement is not clearly correlated with Microtus species of origin, indicating that either spill-over infection or host switching may occur with these viruses. An apparent example exists in the Ohio rodent samples of spill-over of a PH-like virus infection from Microtus species rodents to a deer mouse Peromyscus maniculatus (Pm1047). These viruses have not been associated with HPS cases. The viruses associated with the subfamily Sigmodontinae rodents are highly diverse and are made up of several distinct viruses and lineages in North and South America. All viruses associated with Peromyscus species rodents form a well-supported distinct monophyletic clade (labeled P in Figure 1); these viruses constitute the major cause of HPS cases in North America. Other HPS-associated viruses in this group include Black Creek Canal virus, associated with Sigmodon hispidus. This virus, the cause of a single HPS case, has been genetically detected in cotton rats throughout southern Florida but, so far, nowhere else in the United States. Another genetically distinct virus, Muleshoe virus, has been identified in S. hispidus from the western part of its range (18), but sequences were not available for comparison at the time of our analysis. Caño Delgadito virus, found in S. alstoni in Venezuela (19), appears to be monophyletic with Black Creek Canal viruses. However, bootstrap support for this relationship is low (lower than 50%). Reasonable support is found for the clade containing both these Sigmodon sp.-associated viruses and the Bayou viruses, present in Oryzomys palustris throughout the southeastern United States from the Atlantic coast to Texas (20-22). Bayou viruses have been associated with three HPS cases (20-22). El Moro Canyon virus has been found in numerous harvest mice (Reithrodontomys megalotis) throughout the southwestern United States but has also been found in other rodents (e.g., WA-R2025, in M. montanus), presumably indicating spill-over infections (16,18,23,24). So far, these viruses have not been associated with human disease. The current phylogenetic analysis places these viruses in a distinct supported clade. We analyzed hantaviruses that are also associated with HPS cases in South America and form a well-supported clade that encompasses viruses from Brazil, Argentina, and Paraguay, including the original Juquitiba virus detected in a human autopsy sample from an HPS patient in Brazil in 1993 (25-27). The rodent host for this virus is unknown. Two additional hantavirus lineages have been detected in more recent Brazilian HPS cases (Johnson and Nichol, unpub. data), suggesting that at least three genetically distinct hantaviruses are associated with HPS cases in Brazil. One of these lineages (b9618005) is phylogenetically closer to the Andes virus found in Argentina (28). Andes virus has recently been associated with several HPS cases in Patagonia; its likely host is Oligoryzomys longicaudatus (5,28,29). Finally, Laguna Negra viruses form a well-supported monophyletic lineage. This virus, associated with a large HPS outbreak in the Chaco region of Paraguay, is found in Calomys laucha rodents (10,30). SNV-Like Viruses of Peromyscus Species Rodents We analyzed 229 SNV-like viruses associated with Peromyscus species rodents; they form a well-supported (83%) clade (labeled P in Figure 1; details shown in Figure 2) and are distinct from other Sigmodontinae-associated hantaviruses. These SNV-like viruses include many classic SNVs, which are the major causes of HPS cases throughout the western and central United States and Canada, and are primarily associated with P. maniculatus. These viruses form a distinct, well-supported (78%) clade (labeled S in Figure 2), separate from other SNV-like viruses (Figure 2). Classic SNV 139 bp G2 fragments show up to an 18% nucleotide sequence divergence. Despite a number of exceptions, different genetic variants of SNV are grouped, generally speaking, by geography—an approximate geographic progression is apparent from the north and west toward the south and east, from the top of the tree down toward the node connecting these SNVs (labeled S in Figure 2). For instance, all samples from western Canada, including the Yukon, British Columbia, Alberta, Saskatchewan, and Manitoba are in the upper portion of this clade; two major lineages in California and Nevada (16,31) are also in this clade region. The lower part of the clade is dominated by viruses associated with the original Four Corners outbreak (New Mexico, Colorado, Utah, and Arizona) and other viruses from the Southwest, such as Kansas and Texas. Human HPS cases are represented throughout the SNV clade, indicating that these SNV variants can be associated with HPS illness. In addition to recent samples, 30 SNV-like virus samples from the 1980s were included in the analysis to examine stability of the various SNV genetic lineages and their distribution (labeled H in Figure 2). Only small numbers of nucleotide differences, if any, were observed between old and recent virus sequences from the same geographic areas. The most striking example is the detection of identical viral G2 fragment sequences in rodents captured 12 years apart in New Mexico (Pm434) and Arizona (Pt AZ R29). Similarly, identical viral G2 sequences were found in rodents captured in eastern California in 1983 (our Pm435 and the previously published Sweetwater Canyon sequence [32]) and in human and rodent materials from eastern California and western Nevada sampled 10 or more years later (e.g., Humans CAH19 and NY-H575, and Pm LY-758, 786, and 792). Other examples include 1 of 139 and 2 of 139 nucleotide sequence differences between Washington rodent Pm432 (captured in 1980) and Pm206 and HPS case 0669 (sampled 16 years later), respectively; only 2 of 139 nucleotides are different between Pm428 from southern Oregon and Pm LY-R2302 from northern Nevada, despite capture 12 years apart. These and other data (6,7,32,33) suggest that SNV has been present in North America for a considerable time and has been relatively stably maintained in rodent populations. The next most closely related viruses are those detected in the northeastern United States, referred to as New York virus (34). These viruses have been detected in two human HPS cases and in P. leucopus in New York and Rhode Island (Figure 2). The 139 nucleotide fragments of these viruses have up to 10.1% nucleotide variation, and they differ from classic SNVs by at least 11.5% at the nucleotide level. The next closest group contains viruses associated with several "forest form" subspecies of P. maniculatus throughout the eastern United States and Canada, including the cloudland deer mouse (P. maniculatus nubiterrae), which inhabits the Appalachian mountain region (35). These viruses can also be found in some P. leucopus in this region (e.g., rodent Pl 313 from Pennsylvania). Up to 17.3% nucleotide variation can be seen among the 139 nucleotide fragments of these viruses. The name Monongahela has been suggested for this virus lineage (36), which differs from New York and SN viruses by at least 8.6% and 10.8% nucleotide differences, respectively. Another distinct hantavirus lineage can be seen in P. maniculatus in Tennessee and has been associated with an HPS case (0027) in eastern North Carolina. These viruses are 7.9% different from one another at the nucleotide level for the 139 nucleotide fragment analyzed, and at least 12.2%, 14.4%, and 15.8% different from New York, Monongahela, and SN virus lineages, respectively. Additional distinct virus lineages, recently referred to as Blue River virus (37), can be detected in P. leucopus in Oklahoma (Pl 707), Indiana (e.g., Pl 9436372 and Pl 9436378), and Missouri (e.g., Pl 170). The Oklahoma lineage virus is 10.1%, 10.8%, 15.8% different from the viruses in the Missouri, Indiana, and Tennessee lineages. In addition to identifying the distinct SNV-like viruses and virus genetic lineages throughout North America, our study provides data suggesting the likely site of infection and minimum incubation time for some HPS cases. As reported earlier (2), the HPS case labeled CO H5 was originally described as an Arizona case because the person was residing near Springerville, Arizona, when the illness began. However, the person had been living in Hesperus, Colorado, 11 days before disease onset. The PCR fragment amplified from the case autopsy specimen and from the P. maniculatus trapped at the household in Hesperus matched exactly and differed from those amplified from P. maniculatus in the Arizona location (Figure 2). Similarly, a patient (labeled human 0038) whose symptoms began in Los Angeles, California, had been in the Santa Fe, New Mexico, area 28 to 35 days before illness onset. Analysis of PCR fragments linked the source of infection to New Mexico, rather than to California (Figure 2). Virus and Host Genetic Relationships and Evolution The genetic data we present indicate a broad spectrum of genetic variants of SNV-like viruses throughout North America, associated primarily with Peromyscus rodents. Recent analysis of rodent mitochondrial DNA sequence differences suggests that the different SNV-like virus lineages are primarily associated with different Peromyscus species, and in some cases, with phylogenetically distinct subspecies or mitochondrial DNA haplotypes (Morzunov and Nichol, unpub. data; 37). For instance, the classic SNV and the Monongahela virus lineages are found associated with the "grassland form" and "forest form" of P. maniculatus, respectively (they represent different subspecies and appear phylogenetically distinct with respect to their mitochondrial DNA [Morzunov and Nichol, unpub. data]). The New York virus, and the Blue River virus lineages found in Indiana and Oklahoma, appear associated with genetically distinct P. leucopus populations (37). This pattern likely reflects microadaptation of the virus to the rodent host and not just geographic isolation of the virus variants. This view is supported by the observation that even in areas such as the eastern United States (particularly the Appalachian Mountain region), where P. maniculatus (forest form) and P. leucopus (eastern form) are sympatric and share microhabitat, extensive virus mixing between species is not seen; the Monongahela virus lineage is found predominantly in P. maniculatus, and the New York virus lineage in P. leucopus. Such data suggest that the broad correlation clearly evident between virus evolutionary relationships and those of their primary rodent reservoirs likely exists even at the finer level of closely related species and subspecies. However, the fact that the P. leucopus–associated New York virus appears phylogenetically closer to the P. maniculatus–associated viruses (SN and Monongahela) than to other P. leucopus–associated viruses (Blue River) suggests that this coevolutionary relationship is not absolute and that some species jumping (host-switching) may also have occurred. While the exact phylogenetic relationship of the SNV lineages to Monongahela, New York, and the other P. leucopus virus lineages is not well resolved by using the 139-bp G2 fragment we analyzed, analysis of more complete sequence data strongly supports a similar topology, placing New York virus firmly within the clade of P. maniculatus– borne viruses (37). This evidence, together with significant spill-over infection that sometimes occurs between sympatric rodents, illustrates the complexity of the hantavirus-host interactions. This observation leads into another area of complexity, namely, the definition of distinct hantavirus serotypes or species. In the past, a newly identified arbovirus would be considered a distinct virus or virus serotype if a fourfold or greater two-way difference between this virus and previously recognized closely related viruses was obtained in virus neutralization assays. Despite the obvious biologic limitation (a single amino acid change can allow virus to escape from neutralization), this traditional criterion correlates remarkably well with more recent molecular data. One problem is that hantaviruses are generally difficult to isolate in tissue culture and are frequently noncytopathic, often making plaque assay analysis impractical (Table 1). An attempt to define distinct virus species by using more widely used general criteria for the definition of biologic species is under way. Most defined species could be described as the lowest taxonomic unit that is geographically and ecologically contained, reproductively isolated, phylogenetically distinct, and self-sufficient. The apparent long-term maintenance and coevolution of phylogenetically distinct hantaviruses with different primary rodent reservoir species provides a foundation on which to build a hantavirus species definition. That is, if little host switching has occurred and if instead hantaviruses are associated with specific primary rodent reservoir species for many thousands of years, identification of a hantavirus in a unique primary rodent reservoir species would strongly suggest that in further analyses (e.g., nucleotide and amino acid sequence, cross-neutralization), it will be found to represent a new virus species. Hantaviruses maintained in rodent hosts from different genera (e.g., SNV in Peromyscus species rodents compared with Black Creek Canal virus in Sigmodon species rodents) will clearly meet the broad criteria for separate species status. This view is reinforced by recent data showing that stable reassortant viruses of different SNV genetic lineages can be readily detected in nature (31,38) and in tissue-culture mixed infections (39), but not in virus pairs such as SNV and Black Creek Canal virus (39). Difficulty can arise when trying to determine the species status of viruses maintained within rodent hosts of the same genera or species. So far, SN, New York, Monongahela, and Blue River viruses have been suggested as distinct hantaviruses with independent species names (5,36,37). The genetic analysis we present suggests that, as more hantavirus-infected Peromyscus species samples are analyzed, it is increasingly difficult to draw clear lines separating these virus species. The decision regarding whether to lump these viruses together as SNV-like viruses or to split them into separate species status will require the availability of neutralization data for several representatives of each virus, more detailed identification of the virus-host relationships, and more complete genetic characterization of both viruses and their hosts. Appendix Rodent and HPS Case Materials The newly described nucleotide sequences were derived from rodent materials collected as part of a nationwide survey of rodents for hantavirus antibodies (Ksiazek et al., unpub. data). Most of the human HPS-case blood and tissue autopsy samples were obtained and examined during the original investigation of an HPS outbreak in the Four Corners area of the southwestern United States in 1993 and as part of national surveillance for hantavirus disease throughout the United States from 1993 to 1997. Canadian rodent and HPS case materials were provided by the Laboratory Centre for Disease Control, Canada. Historic rodent samples were obtained from the Division of Biological Materials of the Museum of Southwestern Biology (Albuquerque, NM), University of New Mexico. RNA Extraction, RT-PCR Amplification and Sequencing Total RNA was extracted from human and rodent tissues, blood, or serum (2,10). Because of the hazardous nature of the virus, homogenization of rodent and human autopsy materials and extraction of RNA were performed in a certified class IIb laminar flow biosafety hood in Biosafety Level 3 containment. RNA was extracted from tissue or blood products by using acid guanidinium thiocyanate and phenol-chloroform and purified by using the RNaid Kit (Bio 101, La Jolla, CA). Nested RT-PCR assays were used to amplify DNA products containing a small fragment of the G2 coding region of M segment (2,10). Rodent and human samples were amplified separately, and all manipulations that might result in possible cross-contamination of samples were avoided. PCR products of correct size were sequenced with the same primers used for second-round PCR amplification in conjunction with various generations of sequencing kits available from Applied Biosystems, Inc. (Perkin Elmer, Foster City, CA). Sequences 139 nucleotides in length determined from each PCR product were used in phylogenetic analysis. Oligonucleotide Primer Design Oligonucleotide primers were used to generate DNA fragments from the G2 region of hantavirus M RNA (Table 2). In the initial phase of this project, amplification of hantavirus sequences from autopsy tissues of fatal HPS cases and hantavirus antibody-positive rodents in the southwestern United States used primers designed on the basis of nucleotide sequences conserved among PH and Puumala viruses (2). On the basis of SNV nucleotide sequences derived from these materials, new primers were designed and optimized for detection of SNV-like viruses associated with P. maniculatus (11). As more sequence data became available, additional generations of primers were refined that would detect hantaviruses from other geographic regions of the United States. The development of broadly reactive primers designed to detect hantaviruses associated with subfamily Sigmodontinae rodents (10) has eliminated the effort of amplifying RNA samples with many sets of primers. Table 2. PCR and sequencing primers Basis of primer design Amplicon 1st-round primers (5' to 3') 2nd-round primers (5' to 3') (ref.) size TTTAAGCAATGGTG(C/T)ACTAC(T/A)AC AGAAAGAAATGTGCATTTGC Puumala/ 278 Prospect Hill/ CCATAACACAT(A/T)GCAGC CCTGAACCCCATGC(A/T/C)CCATC Arvicolinae (2) TTTAAGCAATGGTG(C/T)ACTAC(T/A)AC AAGGTAACACAGT(G/C)TCTGGATTC Sin Nombre/ 185 Western CCATAACACAT(A/T)GCAGC GGTTATCACTTAGATC(C/T)TGAAAGG U.S. 1st (2) generation AGAAAGATCTGTGGGTTTGC AAGGTAACACAGT(G/C)TCTGGATTC Sin Nombre/ 185 Western CCTGAACCCCAGGCCCCGT GGTTATCACTTAGATC(C/T)TGAAAGG U.S. 2nd (11) generation TGTGTGTTTGGAGACCCTGG ATGTCAACAAC(A/G)AGTGGGATG Sin Nombre 185 Nevada/ TC(A/G)ATAGATTGTGTATGCA CATGGGTTATCACTTAG(G/A)TC E. California (31) CAGAAAGATCTGCGGGTTTGC CAAGGGAATACTGTCTCTGGATTT Bayou 185 virus/LA/ CCCGAGCCCCATGCACCAT GATTGTCACTCAGATCTTGAAATG East Coast/ S. American (19,20) TGTGAITATCAAGGIAAIAC TGTGAITATCAAGGIAAIAC General 242 Sigmodontinae ACIG(A/T)IGCICCATAICACAT CCCCAIGCICCITCAAT (10) Nucleotide Sequence and Phylogenetic Analyses Compilation, alignment, and comparative nucleotide sequence analysis was carried out by using the Wisconsin Sequence Analysis Package, version 8.1 (Genetics Computer Group, Inc., Madison, WI) on a DEC 3000-500X AXP workstation (Digital Equipment Corp., Maynard, MA). Phylogenetic analysis was performed by maximum parsimony analysis using PAUP version 4.0 d52 (12) on a Power PC 9500. The size and complexity of the dataset prevented the use of branch and bound search method and weighting of the data matrix based on transition:transversion bias. Maximum parsimony analysis of the hantavirus G2 fragment nucleotide sequences was carried out by using the heuristic search option. The initial unweighted analysis showed considerable homoplasy within the dataset. A successive approximations method was used in which characters were weighted by using the maximum value of their rescaled consistency index (12), and the heuristic search repeated. Bootstrap analysis was carried out by 500 replicates of the heuristic search with random resampling of the data. The analysis required several months of computer time; thus, it was not possible to include some recently published additional hantaviruses sequences. The nucleotide sequence dataset (bigtree.nex) we used is available in NEXUS format (compatible with most phylogenetic analysis software packages). A brief description of all samples analyzed is available in EXCEL 2.1 spreadsheet format (bigtree.exl). The phylogenetic tree of Figures 1 and 2 can also be found at the same location on the website. Acknowledgments We thank Ali Khan, Jim Mills, Jamie Childs, John Krebs, Tom Mather, Joe Camp, Fred Jannett, and many others for their efforts in collecting hantavirus infected materials. We also thank Kent Wagoner and Laura Morgan for assistance with graphics and database manipulations. Ms. Monroe is a biologist with the Special Pathogens Branch, Division of Viral and Rickettsial Diseases, CDC. Her research focuses on molecular phylogenetics of hantaviruses and reverse genetic studies of Ebola virus. Address for correspondence: Stuart Nichol, Centers for Disease Control and Prevention, Special Pathogens Branch, Mail Stop G14, 1600 Clifton Rd., N.E., Atlanta, Georgia 30333, USA; fax: 404-639-1118; e-mail: stn1@cdc.gov. References 1. McKee Jr KT, LeDuc JW, Peters CJ. Hantaviruses. In: Belshe RB, editor. Textbook of human virology. 2nd ed. St. Louis (MO): Mosby; 1991. p. 615-32. 2. 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Yanagihara R, Daum CA, Lee P-W, Baek L-J, Amyx HL, Gajdusek DC, et al. Serological survey of Prospect Hill virus infection in indigenous wild rodents in the USA. Trans R Soc Trop Med Hyg 1987;81:42-5. 8. Mills JN, Johnson JM, Ksiazek TG, Ellis BA, Rollin PE, Yates TL, et al. A survey of hantavirus antibody in small-mammal populations in selected United States national parks. Am J Trop Med Hyg 1998;58:525-32. 9. Mills JN, Johnson JM, Ksiazek TG, Ellis BA, Rollin PE, Yates TL, et al. A survey of hantavirus antibody in small-mammal populations in selected United States national parks. Am J Trop Med Hyg 1998;58:525-32. 10. Johnson AM, Bowen MD, Ksiazek TG, Williams RJ, Bryan RT, Mills JN, et al. Laguna Negra virus associated with HPS in western Paraguay and Bolivia. Virology 1997;238:115-27. 11. Spiropoulou CF, Morzunov S, Feldmann H, Sanchez A, Peters CJ, Nichol ST. Genome structure and variability of a virus causing hantavirus pulmonary syndrome. Virology 1994;200:715-23. 12. Swofford DL. PAUP*: phylogenetic analysis using parsimony (*and other methods) [computer program]. Version 4.0. Sinauer, Sunderland, MA; 1998. 13. Hillis DM, Bull JJ. An empirical test of bootstrapping as a method for assessing confidence in phylogenetic analysis. Syst Biol 1993;42:182-92. 14. Lee P, Amyx HL, Yanagihata R, Gajdusek DC, Goldgaber D, Gibbs Jr CJ. Partial characterization of Prospect Hill virus isolated from meadow voles in the United States. J Infect Dis 1985;152:826-9. 15. Parrington MA, Lee PW, Kang CY. Molecular characterization of the Prospect Hill virus M RNA segment: comparison with the M RNA segments of other hantaviruses. J Gen Virol 1991;72:1845-54. 16. Rowe JE, St Jeor SC, Riolo J, Otteson EW, Monroe MC, Ksiazek TG, et al. Coexistence of several novel hantaviruses in rodents indigenous to North America. Virology 1995;213:122-30. 17. Song W, Torrez-Martinez N, Irwin W, Harrison FJ, Davis R, Ascher M, et al. Isla Vista virus: a genetically novel hantavirus of the California vole Microtus californicus. J Gen Virol 1995;76:3195-9. 18. Rawlings JA, Torrez-Martinez N, Neill SU, Moore GM, Hicks BN, Pichuantes S, et al. Cocirculation of multiple hantaviruses in Texas, with characterization of the small (S) genome of a previously undescribed virus of cotton rats (Sigmodon hispidus). Am J Trop Med Hyg 1996;55:672-9. 19. Fulhorst CF, Monroe MC, Salas RA, Duno G, Utrera A, Ksiazek TG, et al. Isolation, characterization, and geographic distribution of Caño Delgadito virus, a newly discovered South American hantavirus (family Bunyaviridae). Virus Res 1997;51:159-71. 20. Morzunov SP, Feldmann H, Spiropoulou CF, Semenova VA, Rollin PE, Ksiazek TG, et al. A newly recognized virus associated with a fatal case of hantavirus pulmonary syndrome in Louisiana. J Virol 1995;69:1980-3. 21. Ksiazek TG, Nichol ST, Mills JN, Groves MG, Wozniak A, McAdams S, et al. Isolation, genetic diversity and geographic distribution of Bayou virus. Am J Trop Med Hyg 1997;57:445-8. 22. Hjelle B, Goade D, Torrez-Martinez N, Lang-Williams M, Kim J, Harris RL, et al. Hantavirus pulmonary syndrome, renal insufficiency and myositis associated with infection by Bayou hantavirus. Clin Infect Dis 1996;23:495-500. 23. Hjelle B, Chavez-Giles F, Torrez-Martinez N, Yates T, Sarisky J, Webb J, et al. Genetic identification of a novel hantavirus of the harvest mouse Reithrodontomys megalotis. J Virol 1994;68:6751-4. 24. Torrez-Martinez N, Song W, Hjelle B. Nucleotide sequence analysis of the M genomic segment of El Moro Canyon hantavirus: antigenic distinction from Four Corners hantavirus. Virology 1995;211:336-8. 25. Nichol ST, Ksiazek TG, Rollin PE, Peters CJ. Hantavirus pulmonary syndrome and newly described hantaviruses in the United States. In: Elliott RM, editor. The Bunyaviridae. New York: Plenum Press; 1996. p. 269-80. 26. da Silva MV, Vasconcelos MJ, Hidalgo NTR, Veiga APR, Canzian M, Marotto PCF, et al. Rev Ins Med Trop Sao Paulo 1997;39:231-4. 27. Vasconcelos MJ, Lima VP, Iversson LB, Rosa MDB, Travassos Da Rosa APA, Travassos Da Rosa ES, et al. Pulmonary syndrome in the rural area of Juquitiba, São Paulo metropolitan area, Brazil. Rev Inst Med Trop Sao Paulo 1997;39:237-8. 28. Lopez N, Padula P, Rossi C, Lazaro ME, Franze-Fernandez MT. Genetic identification of a new hantavirus causing severe pulmonary syndrome in Argentina. Virology 1996;220:223-6. 29. Levis S, Rowe JE, Morzunov S, Enria DA, St Jeor S. New hantavirus causing hantavirus pulmonary syndrome in central Argentina. Lancet 1997;349:998-9. 30. Williams RJ, Bryan RT, Mills JN, Palma RE, Vera I, de Velasquez F, et al. An outbreak of hantavirus pulmonary syndrome in western Paraguay. Am J Trop Med Hyg 1997;57:274-82. 31. Henderson WW, Monroe MC, St Jeor SC, Thayer WP, Rowe JE, Peters CJ, et al. Naturally occurring Sin Nombre virus genetic reassortants. Virology 1995;213:602-10. 32. Nerurkar VR, Song JW, Song KJ, Nagle JW, Hjelle B, Jenison S, et al. Genetic evidence for a hantavirus enzootic in deer mice (Peromyscus maniculatus) captured a decade before the recognition of hantavirus pulmonary syndrome. Virology 1994;204:563-8. 33. Zaki SR, Khan AS, Goodman RA, Armstrong LR, Greer PW, Coffield LM, et al. Retrospective diagnosis of hantavirus pulmonary syndrome, 1978-1993—Implications for emerging infectious diseases. Arch Path Lab Med 1996;120:134-9. 34. Hjelle B, Lee SW, Song W, Torrez-Martinez N, Song JW, Yanagihara R, et al. Molecular linkage of hantavirus pulmonary syndrome to the white-footed mouse, Peromyscus leucopus: genetic characterization of the M genome of New York virus. J Virol 1995;69:8137-41. 35. Hall ER. Mammals of North America. New York: John Wiley and Sons; 1981. 36. Song JW, Baek LJ, Nagle,JW, Schlitter D, Yanagihara R. Genetic and phylogenetic analyses of hantaviral sequences amplified from archival tissues of deer mice (Peromyscus maniculatus nubiterrae) captured in the eastern United States. Arch Virol 1996;141:959-67. 37. Morzunov SP, Rowe JE, Ksiazek TG, Peters CJ, St Jeor SC, Nichol ST. Genetic analysis of the diversity and origin of hantaviruses in Peromyscus leucopus mice in North America. J Virol 1998;72:57-64. 38. Schmaljohn AL, Li D, Negley DL, Bressler DS, Turell MJ, Korch GW, et al. Isolation and initial characterization of a new found hantavirus from California. Virology 1995;206:963-72. 39. Rodriguez LL, Owens JH, Peters CJ, Nichol ST. Genetic reassortment among viruses causing hantavirus pulmonary syndrome. Virology 1998;242:99-106. —————————————————————————————————————————————————————————————————————————— Hantavirus Climatic and Environmental Patterns Associated with Hantavirus Pulmonary Syndrome, Four Corners Region, United States David M. Engelthaler,* David G. Mosley,* James E. Cheek,† Craig E. Levy,* Kenneth K. Komatsu,* Paul Ettestad,‡ Ted Davis,§ Dale T. Tanda,§ Lisa Miller,§ J. Wyatt Frampton,¶ Richard Porter,* and Ralph T. Bryan# *Arizona Department of Health Services, Phoenix, Arizona, USA; †Indian Health Service, Albuquerque, New Mexico, USA; ‡New Mexico Department of Health, Santa Fe, New Mexico, USA; §Colorado Department of Public Health and Environment, Denver, Colorado; ¶Utah Department of Health, Salt Lake City, Utah, USA; and #Centers for Disease Control and Prevention, Albuquerque, New Mexico, USA --------------------------------------------------------------------------- To investigate climatic, spatial, temporal, and environmental patterns associated with hantavirus pulmonary syndrome (HPS) cases in the Four Corners region, we collected exposure site data for HPS cases that occurred in 1993 to 1995. Cases clustered seasonally and temporally by biome type and geographic location, and exposure sites were most often found in pinyon-juniper woodlands, grasslands, and Great Basin desert scrub lands, at elevations of 1,800 m to 2,500 m. Environmental factors (e.g., the dramatic increase in precipitation associated with the 1992 to 1993 El Niño) may indirectly increase the risk for Sin Nombre virus exposure and therefore may be of value in designing disease prevention campaigns. Relationships between environmental characteristics (i.e., climate, biome, elevation, habitat structure, and microhabitat), time, and hantavirus pulmonary syndrome (HPS) cases in the United States have not been systematically evaluated. We describe environmental factors associated with probable exposure sites for all diagnosed HPS cases in the Four Corners region (Arizona, New Mexico, Colorado, Utah) of the United States before 1996 and evaluate the strength of the associations between climate, time, and HPS cases. Case and Exposure Site Identification All cases met the surveillance case definition for HPS, which requires the presence of clinically compatible symptoms and laboratory confirmation (1). Fifty-nine sites in Arizona, New Mexico, Colorado, and Utah were identified as probable exposure sites for HPS cases (sites at which HPS patients were most likely infected with Sin Nombre virus (SNV), according to previously collected data from environmental assessments and individual patient questionnaires [2]) occurring from 1985 to 1995. Rodent trapping and testing data were also used to determine exposure sites, including those identified by linking hantavirus genome sequences between patients and rodents (3,4). Our inability to determine precisely where persons were exposed to HPS may have created a selection bias. Climate Investigations Climate analyses were limited to 52 probable exposure sites of HPS cases (n = 53 cases) (two case-patients were exposed to SNV at the same site); onset of illness was between 1993 and 1995. Climate data for probable exposure sites were collected from the nearest weather station monitored by the National Oceanic and Atmospheric Administration's Western Regional Climate Center in Reno, Nevada. Ten years (1986 to 1995) of monthly precipitation sums and monthly averages of daily ambient temperature were obtained for each station. To be included, sites had to be located within a 30-km radius and a 300-m elevation range from their closest corresponding weather station and without a mountain range between the exposure site and the identified weather station. Twelve sites did not meet these inclusion criteria, six were removed from precipitation analyses, and five were removed from temperature analyses because of missing weather data. Data from 24 weather stations were used for the precipitation (n = 34 exposure sites) and temperature analyses (n = 35 exposure sites). Four weather stations represented multiple exposure sites because they were the closest available weather stations. Twenty-two (85%) exposure sites included in the analyses were within 15 km (mean = 13.2 km, standard deviation [SD] = 7.9 km) and 150 m of elevation (mean = 78.0 m, SD = 79.2 m) of their corresponding weather station. Data were analyzed with the SPSS and Epi-Info statistical software programs (5,6). The use of climatic data collected from weather stations as far as 30 km away from probable exposure sites, with as much as a 300-m difference in elevation, may have created an ascertainment bias. Weather patterns, however, generally cover large regions. In addition, a withdrawal bias may have been introduced by eliminating exposure sites and weather stations that did not meet the inclusion criteria. Precipitation Analysis Precipitation totals from the 24 identified weather stations for each of the 48 months during 1992 through 1995 were compared with the corresponding calendar month's mean precipitation total for 1986 to 1991. Each month's mean precipitation difference was plotted against the number of HPS cases with onset of symptoms during that month for cases between 1993 and 1995. The Wilcoxon matched-pairs signed rank test was used to test the statistical significance of precipitation during two periods of substantial departure from normal precipitation patterns (i.e., 6-year mean sum precipitation data for the same calendar months). These weather stations, representing the 1993 to 1995 cases, reported above average (p < 0.01) precipitation totals during December 1992 through March 1993 and below average (p < 0.01) precipitation totals during June 1993 through July 1993 (Figure 1). By using a Spearman's correlation, we found a negative correlation between the number of cases per month (onset [Fig 1] date) and the number of months after Figure 1. Mean difference in monthly the 1992 to 1993 El Niño weather precipitation and temperature pattern (r(sub s)= -0.70; p < 0.01) between month of interest and 6-year mean (1986–1991) at the study sites (Figure 2). For this portion of the and number of cases by month, precipitation analysis, the months of 1993–1995. onset were used for all 53 cases between 1993 and 1995. The end of the 1992 to 1993 El Niño was described as the month (March 1993) in which the abnormally high precipitation totals [Fig 2] for November 1992 through March 1993 Figure 2. Correlation of hantavirus returned to normal, on the basis of pulmonary syndrome (HPS) cases per the data in Figure 1. month and number of months from end of El Niño period (n = 53 cases). The 1993 outbreak of HPS in the Four Corners region followed a dramatic increase in precipitation associated with the 1992 to 1993 El Niño phenomenon and peaked in the middle of a drought. Because the 1992 to 1993 El Niño resulted in an abundance of rodent food resources (e.g., vegetation and insects) and a 20-fold rodent population increase over the previous year at the Sevilleta National Wildlife Refuge in central New Mexico, increased rainfall from El Niño was associated with the 1993 HPS outbreak (7). A similar pattern of above average rainfall followed by drought was observed preceding an outbreak of HPS in western Paraguay in 1995 to 1996 (8). Our study shows that the number of HPS cases per month in the Four Corners region during 1993 to 1995 was negatively correlated with the number of months after the 1992 to 1993 El Niño. The data suggest that the association between the 1992 to 1993 El Niño and the number of HPS cases in the Southwest may have lasted for as long as 2 years. The association between El Niño and the HPS outbreak is probably complex. The above average precipitation during the winter and spring of 1992 to 1993 may have increased rodent populations and thereby increased the likelihood of more rodent-to-rodent contact, rodent-to-human contact, and viral transmission, thereby resulting in the large number of cases in 1993 and 1994. In addition, as rodent populations surpassed the carrying capacity of their local environments and precipitation plummeted, available food sources may have been depleted, resulting in rodent population stress. Increased stress likely increased rodent-to-rodent contact, as rodents competed for food and water, and increased rodent-to-human contact, as rodents moved into new, potentially less stressful environments, such as homes and peridomestic structures. During 1995, no cases occurred in the original outbreak area near the Arizona-New Mexico border and in western Colorado, possibly because the effect of the 1992 to 1993 El Niño had dissipated. Preliminary data from longitudinal trapping studies in the Southwest suggest that the relative rodent densities decreased to normal levels in 1995 (J. Mills, T. Yates, pers. comm.). Hantavirus infection rates in rodents dramatically decreased at case sites in Arizona and Colorado 3 years after the outbreak (9). Temperature Analysis Monthly ambient temperature means from the 24 identified weather stations for each of the 48 months during 1992 through 1995 were compared to the corresponding calendar month's ambient temperature mean for 1986 to 1991. The mean difference was plotted against the number of HPS case onsets per month for cases occurring between 1993 and 1995. With all 24 weather stations included, the Wilcoxon matched-pairs signed rank test was used to test the statistical significance of two periods with substantial departures from normal temperature patterns (i.e., 6-year mean daily temperature averages for the same calendar months): 1) November 1992 through December 1992 and 2) January 1993. The 24 weather stations reported below average (p < 0.01) temperatures during November 1992 through December 1992, which corresponded with the onset of El Niño (Figure 1). Conversely, the mean temperature for January 1993 was above average (p < 0.01). While interpreting variations in precipitation totals is fairly straightforward, interpreting variations in ambient temperature is far more difficult. Whether the lower-than-average ambient temperature during November 1992 through December 1992 affected rodent population dynamics is unknown; however, the higher-than-average temperatures during January 1993 may have promoted rodent survival during what is normally the coldest month of the year. (Figure 1 indicates that the 24 weather stations reported the most substantial mean temperature extremes in 1995.) To examine the relationship between ambient temperature and month of exposure for HPS cases, we compared the distribution of daily ambient temperatures of probable month (defined as the month in which onset date minus 14 days [hypothesized mean incubation period] occurred) of exposure to daily ambient temperatures during the other 11 months of the same calendar year. Exposure months had higher ambient temperatures and a smaller temperature range than the other months in the same calendar year. The mean temperature for the probable exposure months was 15°C (SD = 7°C; range = 0°C - 24°C) (Figure 3), and the mean throughout the rest of the year was 11°C (SD = 9°C; range = -7°C - 36°C). Spatial, Temporal, and Environmental Investigations Spatial and environmental data were [Fig 3] collected for 59 of the 64 known HPS Figure 3. Distribution of monthly cases that occurred before 1996 in the ambient temperatures for months Four Corners region: Arizona (20 estimated as the month SNV sites), New Mexico (25 sites), exposure occurred and the 11 Colorado (7 sites), and Utah (7 nonexposure months for the same sites). For five cases (in 1959, 1975, calendar year (n = 35 cases and 24 1984, 1985, and 1993), the exposure weather stations). site was not known. Habitat data (e.g., dominant biome type as described by Brown [10] within 200 m) were collected at each of the 59 probable exposure sites. Elevations were taken from U.S. Geological Service quadrant maps. All analyses were conducted with the SPSS and Epi-Info statistical software programs (5,6). Spatial Analysis Figures 4, 5, and 6 display the spatial and temporal distribution of probable hantavirus exposure sites in the Four Corners region, from 1993 through 1995 (n = 52). In 1993, most case-patients were exposed in northeastern Arizona and northwestern New Mexico; in 1994, HPS case-patients were exposed in all four states, including two new areas, central Utah and southern Arizona; and in 1995, HPS case-patients were exposed only in a small geographic area near the border of southcentral Colorado and northcentral New Mexico (on the eastern slope of the Rocky Mountains). From 1993 through 1995, HPS cases in the Four Corners region shifted geographically. The apparent [Fig 4] spatial and temporal movement of Figure 4. Hantavirus pulmonary HPS foci in this region mirrors the syndrome cases in the Four Corners "focal nidality" exhibited by region, by probable exposure site hemorrhagic fever with renal location, 1993–1995 (n = 53 cases and syndrome (HFRS), the Eurasian 52 exposure sites). hantaviral manifestation (11-15). HFRS cases can occur sporadically throughout the year, but outbreaks occur seasonally (12,16). Seasonal prevalence varies by locality and meteorologic and climatic conditions that favor activity of rodents associated with viruses causing HFRS (12). Niklasson et al. (17) hypothesize that large outbreaks of HFRS occur in geographic "hot spots," which may depend on certain ecologic characteristics correlating with rodent populations, e.g., rodent habitats and environmental or meteorologic conditions (17). In the Four Corners states, seroprevalence of hantaviruses in rodents displays a similar focality (18). Our data suggest that HPS cases may have occurred in similar hot spots in this region between 1993 and 1995. Temporal Analysis The 1993 to 1995 HPS onset months were unevenly distributed (Kolmogrov-Smirnov test, p = 0.01), displaying a spring-summer seasonality, as has been reported previously (2,3,15) (Figure 6). While cases occurred throughout all seasons, probable exposures occurred during the time of year when the monthly mean ambient temperature was 0°C to 24°C. [Fig 5] Figure 5. Hantavirus pulmonary syndrome cases in the Four Corners region by state, 1993–1995 (n = 53 cases and 52 exposure sites). [Fig 6] Figure 6. Hantavirus pulmonary cases in the Four Corners region, by month of onset, 1993–1995 (n = 53 cases and 52 exposure sites). The seasonality of HPS cases varies by location, elevation, and biome (Figures 4, 5, and 7). The clustering of all four Sonoran Desert HPS cases during the late winter and early spring and the two montane conifer cases during the late spring and summer, when the mean temperatures for both biomes are mild (12°C - 21°C), are the best examples of this trend. Most grassland, Great Basin scrub, pinyon-juniper, and montane conifer cases occurred during spring and summer (Figure 7). Environmental Analysis Probable exposure sites occurred in seven biomes, most often in pinyon-juniper woodland, grassland, and Great Basin desert scrub biomes (33.9%, 28.8%, and 23.7%, respectively) (Table). Approximately 66% (39/59) of case-patients were exposed at elevations of 1,800 m to 2,500 m; none was exposed at elevations higher than 2,500 m (Table). This description may reflect the typical biomes in which the 1993 outbreak took place but also holds for cases in Utah and Colorado. Mills et al. (18) reported the same habitat and elevation for sites with the highest Peromyscus densities (as well as the highest rodent SNV antibody prevalence) in the Four Corners states. Unpublished case-control data (J. Cheek and R. Bryan) show that significantly more HPS cases occurred in Great Basin pinyon-juniper than in any other biome on the Navajo reservation in the Southwest; the data also show that HPS cases are more likely at higher elevations. Environmental conditions surrounding HPS exposure sites appear similar to those surrounding plague exposure sites on the Navajo reservation (K. Gage, R. Enscore, unpub. data). Probable exposure sites were most frequently found in rural areas, with five or fewer homes within a 1-km radius (64.4%); most exposure sites [Fig 1] (88.1%) were found in areas with 25 or Figure 7. Number of hantavirus fewer homes. An almost equal number of pulmonary syndrome cases occurring exposure sites (44.1%) was more than 2 in specific biomes by season of km away from a permanent water source onset (1993–1995) (n = 53 cases). as exposure sites (42.4%) within 1 km of a permanent water source. Overall, 104 plant species belonging to 72 genera were identified at exposure sites. The three most common dominant herbaceous species were Artemisia tridentata (big sagebrush), Gutierrezia sarothrae (snakeweed), and Bromus tectorum (downy chess grass); and the three most common dominant overstory and understory species were Juniperus monosperma (one-seed juniper), J. osteosperma (Utah juniper), and Pinus edulis (two-needle pinyon pine). For 21 of 59 sites with 5% slope or greater, more than twice as many had a northward aspect (9 of 21) than any other aspect (south = 4 of 21; east = 4 of 21; and west = 4 of 21). While the habitat may have changed in minor ways between the time of exposure and data collection, no major habitat changes (e.g., fire, timber harvesting) were identified. Because these environmental findings are descriptive, caution should be used when drawing conclusions on the basis of these data. Conclusions Table. Frequency of probable Sin Nombre virus exposure sites in selected biomes and elevation Previous studies have ranges examined behavior and environmental risk ------------------------------------------------- factors for the Frequency acquisition of HPS Biome type (n = 59) % (19-22). Certain behavior ------------------------------------------------- and occupations may increase the likelihood Sonoran Desert 4 6.8 of exposure to the Chihuahuan Desert 1 1.7 excreta of infected Great Basin Desert scrub 14 23.7 rodents; however, the overall level of risk Great Basin and plains grassland 17 28.8 seems low (21,22). Great Basin pinyon-juniper 20 33.9 Environmental risk factors may be divided Riparian woodland 1 1.7 into endemic and Montane conifer forest 2 3.4 epidemic. Endemic Elevation range environmental risk factors are tied to the ------------------------------------------------- static habitat structure 0 - 619 3 5.1 of a geographic area. For 620 - 1,239 1 1.7 example, the habitat of the Four Corners region 1,240 - 1,859 13 22.0 allows population levels 1,860 - 2,480 42 1.2 of Peromyscus maniculatus great enough to maintain ------------------------------------------------- SNV transmission (18,23). Some level of risk will likely always exist for exposure to SNV in the region; however, the level seems low (9). Epidemic environmental risk factors, also tied to the static habitat structure of a geographic area (i.e., they can only occur in disease-endemic areas), are primarily dynamic events associated with large-scale environmental changes of limited duration. For example, the precipitation pattern during the 1992 to 1993 El Niño, associated with increased rodent populations in the Southwest (7) and consequently with the 1993 outbreak of HPS in the Four Corners region, greatly increased the level of risk for human SNV infection in the outbreak area. By 1995, rodent populations (J. Mills, T. Yates, pers. comm.) and the number of HPS cases in this area dramatically decreased, and the level of risk for human SNV infection may have returned to endemic levels in 1995. Early recognition of these endemic and epidemic environmental risk factors might allow public health agencies to predict where and when the next HPS outbreak may occur and thus to effectively target prevention efforts. While this report does not address data related to cases after 1995, the 1997 to 1998 El Niño may not have displayed the same precipitation pattern in the Four Corners region as the 1992 to 1993 El Niño. Limited data from Arizona's primary weather station for northern Arizona indicate 4.70 cm of precipitation in December 1997—the 98-year mean (1899–1997) for December is 4.88 cm (Office of the State Climatologist—Arizona State University, pers. comm.). In December 1992, the same station reported 17.22 cm of precipitation (Office of the State Climatologist—Arizona State University, pers. comm.). Future studies should examine and compare the two precipitation events, their lasting effects on rodent populations, and the subsequent effects on hantavirus epidemiology. Acknowledgments We thank Rosalyn Curtis, Herman Shorty, Rusty Enscore, Dave Tibbs, Chuck Freeman, Benny Joe, Bobby Villines, Eric Faist, Charlie Irland, Tim Doyle, and Dorothy Miller for their assistance in data collection and Charles Calisher, T. Michael Fink, Nicole Rossi, Mira Leslie, and Thomas G. Engelthaler for their critical review and comments. This study was supported in part by CDC cooperative agreements with Arizona, Colorado, New Mexico, and Utah. David Engelthaler is a visiting fellow, Division of Vector-Borne Infectious Diseases, Centers for Disease Control and Prevention, Fort Collins, CO. Mr. Engelthaler is involved in plague research. His primary research interest is pathogen-vector relationships. Address for correspondence: David G. Mosley, Bureau of Epidemiology and Disease Control, Arizona Department of Health Services, 3815 North Black Canyon Highway, Phoenix, Arizona 85015, USA; fax: 602-263-4956; e-mail: dmosley@hs.state.az.us. References 1. Centers for Disease Control and Prevention. Case definitions for infectious conditions under public health surveillance. MMWR Morb Mortal Wkly Rep 1997;46(No.RR-10):16. 2. Khan AS, Khabbaz RF, Armstrong LR, Holman RC, Bauer SP, Graber J, et al. Hantavirus pulmonary syndrome: the first 100 U.S. cases. J Infect Dis 1996;173:1297-303. 3. Hjelle B, Torrez-Martinez N, Koster FT, Jay M, Ascher MS, Brown T, et al. Epidemiologic linkage of rodent and human hantavirus genomic sequences in case investigations of hantavirus pulmonary syndrome. J Infect Dis 1996;173:781-6. 4. Nichol ST, Spiropoulou CF, Morozunov S, Rollin PE, Ksiazek TG, Feldman, et al. Genetic identification of a novel hantavirus associated with an outbreak of acute respiratory illness in the southwestern United States. Science 1993;262:615-8. 5. SPSS for Windows [computer program]. Version 6.1. Chicago: SPSS, Inc.; 1994. 6. Dean AG, Dean JA, Coulombier D, Brendel KA, Smith DC, Burton AH, et al. Epi Info [computer program]. Version 6. A word processing, database, and statistics program for epidemiology on microcomputers. Atlanta (GA): Centers for Disease Control and Prevention; 1994. 7. Parmenter RR, Brunt JW, Moore DI, Ernest S. The hantavirus epidemic in the Southwest: rodent population dynamics and the implications for transmission of hantavirus-associated adult respiratory distress syndrome (HARDS) in the Four Corners region. Department of Biology, University of New Mexico, Albuquerque, New Mexico; 1993. Sevilleta Long-Term Ecological Research Program (LTER); Publication No.: 41. 8. Williams RJ, Bryan RT, Mills JN, Palma RE, Vera I, De Velasquez F, et al. An outbreak of hantavirus pulmonary syndrome in western Paraguay. Am J Trop Med Hyg 1997;57:274-82. 9. Engelthaler DE, Levy CL, Fink TM, Tanda D, T Davis. Decrease in seroprevalence of antibodies to hantavirus in rodents from 1993-1994 hantavirus pulmonary syndrome cases. Am J Trop Med Hyg 1998;58:737-8. 10. Brown DE. Biotic communities: southwestern United States and northwestern Mexico. Salt Lake City: University of Utah Press; 1994. li. 49-221. 11. McKee KT Jr, LeDuc JW, Peters CJ. Hantaviruses. In: Belshe RB, editor. Textbook of human virology. 2nd ed. St Louis (MO): Mosby Year Book; 1991. li. 615-32. 12. Mayer CF. Epidemic hemorrhagic fever of the Far East, or epidemic hemorrhagic nephroso-nephritis. The Military Surgeon 1952;110:276-85. 13. Gauld RL, Craig JP. Epidemiological pattern of localized outbreaks of epidemic hemorrhagic fever. American Journal of Hygiene 1954;59:32-8. 14. Pon E, McKee KT Jr, Diniega BM, Merrell B, Corwin A, Ksiazek TG. Outbreak of hemorrhagic fever with renal syndrome among U.S. Marines in Korea. Am J Trop Med Hyg 1990;42:612-9. 15. Chapman LE, Khabbaz RF. Etiology and epidemiology of the Four Corners hantavirus outbreak. Infectious Agents and Disease 1994;3:234-44. 16. World Health Organization. Haemorrhagic fever with renal syndrome: memorandum from a WHO meeting. Bull World Health Organ 1983;61:269-75. 17. Niklasson B, Jonsson M, Widegren I, Persson K, LeDuc J. A study of nephropathia epidemica among military personnel in Sweden. Res Virol 1992;143:211-4. 18. Mills JN, Ksiazek TG, Ellis BA, Rollin P, Nichol ST, Yates TL, et al. Patterns of association with host and habitat: antibody reactive with Sin Nombre virus in small mammals in the major biotic communities in the southwestern United States. Am J Trop Med Hyg 1997;56:273-84. 19. Zietz PS, Butler JC, Cheek JE, Samuel MC, Childs JE, Shands LA, et al. A case-control study of hantavirus pulmonary syndrome during an outbreak in the southwestern United States. J Infect Dis 1995;171:864-70. 20. Childs JE, Krebs JW, Ksiazek TG, Maupin GO, Gage K, Rollin PE, et al. A house-hold based, case-control study of environmental factors associated with hantavirus pulmonary syndrome in the southwestern United States. Am J Trop Med Hyg 1995;52:393-7. 21. Armstrong LR, Khabbaz RH, Childs JE, Rollin PE, Martin ML, Clarke M, et al. Occupational exposure to hantavirus in mammalogists and rodent workers [abstract]. Am J Trop Med Hyg 1994;51:94. 22. Zietz PS, Graber JM, Voorhees RA, Kioski C, Shands LA, Ksiazek TG, et al. Assessment of occupational risk for hantavirus infection in Arizona and New Mexico. J Occup Environ Med 1997;39:463-7. 23. Childs JE, Ksiazek TG, Spiropoulou CF, Krebs JW, Morzunov S, Maupin GO, et al. Serologic and genetic identification of Peromyscus maniculatus as the primary rodent reservoir for a new hantavirus in the southwestern United States. J Infect Dis 1994;169:1271-80. —————————————————————————————————————————————————————————————————————————— Hantavirus The following articles on hantavirus contain preliminary results of 3 years of longitudinal mark-release-recapture studies in the southwestern United States, provide an analysis of the sensitivity of the field techniques and statistical analyses for detecting changes in rodent population densities, and discuss the role of longitudinal studies in understanding reservoir host ecology as it relates to human disease. Although the data presented in the articles are overall standardized, methods used to extract information vary by research institution. Long-Term Studies of Hantavirus Reservoir Populations in the Southwestern United States: Rationale, Potential, and Methods James N. Mills, Terry L. Yates, Thomas G. Ksiazek, C.J. Peters, and James E. Childs Centers for Disease Control and Prevention, Atlanta, Georgia, USA --------------------------------------------------------------------------- Hantaviruses are rodent-borne zoonotic agents that cause hemorrhagic fever with renal syndrome in Asia and Europe and hantavirus pulmonary syndrome (HPS) in North and South America. The epidemiology of human diseases caused by these viruses is tied to the ecology of the rodent hosts, and effective control and prevention relies on a thorough understanding of host ecology. After the 1993 HPS outbreak in the southwestern United States, the Centers for Disease Control and Prevention initiated long-term studies of the temporal dynamics of hantavirus infection in host populations. These studies, which used mark-recapture techniques on 24 trapping webs at nine sites in the southwestern United States, were designed to monitor changes in reservoir population densities and in the prevalence and incidence of infection; quantify environmental factors associated with these changes; and when linked to surveillance databases for HPS, lead to predictive models of human risk to be used in the design and implementation of control and prevention measures for human hantavirus disease. Hantaviruses (genus Hantavirus, family Bunyaviridae) are rodent-borne zoonotic agents that cause mild to severe hemorrhagic fevers throughout most of Europe, Asia, and the Americas. The epidemiology of these hemorrhagic fevers is largely defined by the distribution and ecology of the rodent hosts of the viruses. Hantaviruses have been identified at a dramatically increased rate in recent years; some 30 hantaviruses are now recognized throughout the world (1,2). With very few exceptions, each virus is associated with a single primary rodent host of the family Muridae. The rodent, in which the virus establishes a chronic infection, sheds infectious virus into the environment in urine, feces, and saliva (3,4); these characteristics are key to the transmission of the virus, both to humans (most frequently by inhalation of infectious aerosols [5]) and among rodents (frequently by aggressive encounters and biting [6,7]). Human diseases due to hantaviruses, which have been recognized at least since World War I and probably occurred much earlier (8), were unknown in the Americas until recently. Although Rattus norvegicus infected with Seoul virus is common in many cities throughout the Americas (9), human disease associated with a rat-borne hantavirus was not documented in a U.S. city until 1994 (10). Prospect Hill virus, an indigenous North American hantavirus, was isolated from the meadow vole (Microtus pennsylvanicus) as early as 1982 (11) but has never been associated with human disease. Since 1993, when hantavirus pulmonary syndrome (HPS) was recognized and its etiologic agent, Sin Nombre virus (SNV), was isolated and associated with the deer mouse (Peromyscus maniculatus) (12,13), at least 20 New-World hantaviruses, all associated with the same group of indigenous American rodents (family Muridae, subfamily Sigmodontinae) have been described, and HPS has been diagnosed from Canada to Patagonia. The severity (approximately 50% death rate) and wide geographic distribution of this rodent-borne zoonotic disease has prompted intensive collaboration between public health investigators and ecologists to elucidate the ecologic and epizootiologic features of infection in host populations and the factors that lead to human infection. Because no specific treatment is yet available, prevention measures are essential in decreasing HPS-related illness and death. Developing effective preventive measures requires a detailed knowledge of the ecology and epizootiology of hantavirus infection in reservoir populations and the specific situations and mechanisms that result in the transfer of hantaviruses from hosts to humans. Reservoir Studies Reservoir studies, whose role in understanding, controlling, and preventing human disease has been reviewed, have resulted in a series of research goals that may facilitate the collection of data concerning reservoir ecology, a subject pertinent to human health (14). The first goal is to identify the reservoir host; others are a) to determine the area in which the disease may be endemic by identifying the geographic range of the host and the range of infection by the pathogen within the host range; b) to more precisely define relative risk to humans by determining the distribution of the host and pathogen among distinct habitats regionally; c) to investigate potential mechanisms of pathogen transmission within host populations by conducting cross-sectional surveys to define the prevalence of infection among various subpopulations of the host (e.g., male versus female, juvenile versus adult); d) to conduct long-term prospective studies to explain the temporal patterns of infection in host populations; and e) to integrate the data from these studies in a predictive model that will allow early identification of specific times and places where conditions can increase rodent populations or infection in rodent populations and elevate the risk for human disease. This model could be used to minimize the incidence of human disease through public education, habitat modification, or reservoir control. Investigations of HPS cases in the United States have resulted not only in studies of the deer mouse and SNV but also in the identification of three additional host-virus relationships that maintain hantaviruses responsible for human disease (New York virus carried by the white-footed mouse, Peromyscus leucopus [Figure 1] [15]; Black Creek Canal virus carried by the cotton rat, Sigmodon hispidus [16]; and Bayou virus carried by the rice rat, Oryzomys palustris [17]). Most HPS cases in the United States have been caused by SNV. Intensive studies of deer mouse populations have addressed most of the proposed goals. One of the most common and most extensively studied [fig 1] small mammals in North America, the Figure 1. White-footed mouse deer mouse has a well-known (Peromyscus leucopus). Photo by R.B. geographic distribution (18). Forbes, Mammal Image Library of the Antibody screening of deer mouse American Society of Mammalogists. populations throughout North America has provided evidence of SNV infection throughout most of the species' range (Ksiazek et al., unpub. data). Regional studies have shown differences in the prevalence of hantavirus infection among deer mouse populations in different habitats and helped define the varying disease risk to humans in these habitats (7). Finally, studies of the age- or size-specific prevalence of hantavirus infection among reservoir populations have shown that SNV, and other hantaviruses, are transmitted horizontally within reservoir populations, and that one important specific mechanism of transfer may be aggressive encounters and bites, most frequently among male animals (6,7,19). Although these cross-sectional studies have increased our understanding of host-virus ecology as it relates to human disease, they have not explained the temporal dynamics of host-virus ecology nor have they identified the environmental factors associated with these dynamics; only long-term prospective studies can provide this additional information. Long-Term Studies Long-term studies, widely regarded by ecologists as indispensable for understanding the temporal dynamics of vertebrate communities (20), are especially useful for assessing the effects of rare events (e.g., El Niño southern oscillation) and for detecting and observing processes that unfold slowly in communities or populations (e.g., establishment or disappearance of a reservoir species from part of its range; changes in reservoir population density; changes in community composition; introduction or extinction of a pathogen in a specific host population; and changes in the incidence or prevalence of infection within the host population). Long-term studies of reservoir populations have helped elucidate the temporal dynamics of hantavirus infection in host populations for Seoul and Prospect Hill viruses (21) and identify characteristics of reservoir ecology associated with outbreaks of human disease. The numbers of cases of hemorrhagic fever with renal syndrome due to Puumala virus in Scandinavia and Argentine hemorrhagic fever due to Junín virus (an arenavirus with many epidemiologic similarities to hantaviruses) were correlated with cyclic changes in the density of reservoir host populations (22,23). Increases in population density are associated with improved reproductive success and survivorship that may be due to improved habitat. Changes in the environment may be associated with favorable weather patterns, accelerated vegetation growth, and availability of plant and small-animal foods (14,23). The 1993 HPS outbreak in the southwestern United States may have resulted from improvements in the quality of deer mouse habitat caused by the 1991-92 El Niño southern oscillation (24). When the environmental variables associated with increasing reservoir population densities are identified and quantified, a key component of a predictive model of human risk will be in place. Despite their importance and utility, long-term studies of reservoir populations associated with zoonotic agents are rare. By definition, they require stable funding for many years, they are labor intensive, expensive, and may not produce significant results in the short term. The periodic shifts in environmental conditions that change host populations and increase risk for human disease may take many years. The most common method for conducting long-term studies of small-mammal populations is the mark-release-recapture (MRR) technique. Animals trapped live on permanent trapping plots are measured, sampled (blood or oral swab), identified with a permanent mark or number, and released at the exact site of capture. The trapping plots are operated at predetermined intervals for several days. Animals recaptured in subsequent trappings are measured and sampled again so that changes in numbers of animals, body growth rates, movement, reproductive condition, and infection status can be monitored. Environmental variables such as weather conditions and vegetative cover also may be monitored on the trapping plots. Control plots, where invasive procedures are minimized, may be necessary for determining or correcting for the influence of sampling methods on animal survival or population size. After the 1993 HPS outbreak, the Centers for Disease Control and Prevention (CDC) established a network of hantavirus and rodent monitoring sites in the southwestern United States to 1) monitor and quantify the seasonal and year-to-year changes in host population density and the prevalence and incidence of hantavirus infection, 2) identify and quantify the biotic and abiotic environmental factors associated with and likely influencing these dynamics, 3) identify mechanisms of virus transmission within reservoir populations, and 4) identify and measure any effects of infection on individuals and populations of the host. The studies should lead to predictive models of human risk for hantavirus infection and should facilitate prevention and control of human hantavirus disease. Study Sites Arizona, Colorado, and New Mexico were chosen as the general study area because of their high numbers [fig 2] of HPS cases at the time the study Figure 2. Geographic locations of was being designed. In addition, nine sites where mark-release- four sigmodontine rodent species recapture webs are being identified as hantavirus reservoirs operated to study rodent reservoirs inhabit at least parts of the of hantaviruses in a three- state three-state area (P. maniculatus, P. area of the southwestern United boylii, Reithrodontomys megalotis, States. PCMS=Pinyon Canyon Maneuver and S. hispidus). Longitudinal MRR Site (U.S. Army). studies are being conducted on 24 trapping webs at nine sites in the three states (Figure 2): 10 webs at four sites in New Mexico, operated by the University of New Mexico; six webs at three sites in Colorado, operated by Colorado State University; four webs at one site in northern Arizona, operated by Yavapai College; and four webs at one site in southern Arizona, operated by the University of Arizona. Site selection criteria included presence of populations of Peromyscus spp. and evidence of infection by SNV or related viruses in these populations. The location of each trapping web was fixed precisely by global positioning system technology. A sketch of each trapping web site, including a description of the vegetation, was prepared. Placing Permanent Trapping Webs Small-mammal populations were monitored through the use of permanent trapping webs (25) (Figure 3A). Each web covered 3.14 ha and contained 12 100-m transects radiating from a central point and resembling the spokes of a wheel (Figure 3B). Each web contained 148 Sherman (8 x 9 x 23 cm; H.B. Sherman Trap Company, Tallahassee, FL) and 24 Tomahawk (14 x 14 x 40 cm; Tomahawk Live Trap Company, Tomahawk, WI) live-capture traps, at 12 trap stations along each radiating spoke. The first four trap stations were at 5-m intervals and the remaining eight at 10-m intervals. Four Sherman traps were placed around the central point. In addition to a Sherman trap, one Tomahawk trap was placed at trap stations 7 and 12 in each radiating arm (Figure 3B). Two to four webs were located at each sampling site. At least one web at each site was designated a control web. At this web, rodent populations were monitored but not sampled by blood and oral swab collection so that the effects of sampling on small-mammal survivorship could be assessed. At the remaining webs, virus activity in small-mammal populations was monitored through monthly blood and oral swab samples from captured animals. After the second year of the study, the purpose of the control webs was achieved, so sampling of captured small mammals from these webs was initiated (25,26). Trapping Schedules All trapping web sites were visited monthly, except those in Colorado, [fig 3] which were visited every 6 weeks, Figure 3. A. Characteristics of as weather permitted. Webs were landscape and vegetation near Fort operated for 3 consecutive nights on Lewis trapping web "A," southwestern each trapping occasion, generally Colorado. Photo courtesy of C. coinciding with the new moon. Traps Calisher. B. Schematic representation were set out in the evening of the of a trapping web showing the relative first day and baited with peanut locations of the 148 trap stations. butter and rolled oats, cracked Small circles indicate the location of corn, or mixed grain. In cold one Sherman trap, larger circles, one weather, cotton or polyester Sherman plus one Tomahawk trap. fiberfill was placed in the traps to Diameter of the web was 200 m. After provide nesting material and reduce Parmenter et al. (25). trap-associated deaths. Captured rodents were collected, transported, and sampled according to standardized procedures (27,28). Briefly, traps were checked for captures early each morning. Investigators wearing rubber gloves collected the traps containing captured animals, labeled them with the web and trap station number, and placed them in double plastic bags for transport to a centralized outdoor processing station. Before opening the bags containing captured small mammals, investigators donned protective clothing, including latex gloves, disposable surgeon's gowns, and respirators fitted with HEPA filters. Each captured animal was processed individually. The animal was first shaken from the trap into a plastic bag containing cotton or gauze soaked with inhalant anesthesia (methoxyflurane, Pitman-Moore, Mundelein, IL; or isoflurane, Abbott Laboratories, North Chicago, IL). To prevent potential cross-infection between animals, each was anesthetized in a clean plastic bag, and the anesthesia-soaked cotton was contained in a tea strainer that allows diffusion of the anesthesia, yet between animals can be wiped with a disinfectant. In one case, investigators used a specially adapted "nose cone" for anesthesia (Abbott et al., this issue, pp. 102-112). After being anesthetized, the animal was removed from the bag and placed on a clean surface. A standardized form was used at all trapping sites to collect the following data (28): unique capture number; date of capture; exact location of capture on the trapping web (trap station number); ear tag number; fate (first capture, recapture [different trapping session], or repeater [within same 3-day trapping session]); species; age (juvenile, subadult, or adult); mass; lengths of tail plus body, tail only, ear, and right hind foot; reproductive status including position of the testes (scrotal or abdominal) for males and condition of the vagina (closed or perforate) and description of the nipples (enlarged or small, lactating or not) for females; and the presence or absence of scars or wounds. For animals from the sampling webs, oral swabs were taken with Dacron-tipped applicators cut with scissors at the level of the Dacron and inserted into 0.5 ml of virus medium (phosphate-buffered saline containing 20% fetal bovine serum, 2% penicillin and streptomycin, and 0.1% Fungizone) in a 2-ml cryovial. Approximately five drops of whole blood were collected into a second cryovial by a capillary tube inserted into the retro-orbital capillary plexus. Whole blood and oral swab samples were immediately placed in liquid nitrogen or on dry ice until transferred to -70°C freezers for storage. Animals recaptured on days 2 or 3 of the trapping session were not bled a second time to avoid trauma. Animals newly captured were marked with a uniquely numbered ear tag (some smaller animals were marked by toe clipping). The animal was then replaced in the original trap or in a clean, ventilated one-quart, screw-capped jar, and was allowed to recover fully from the effects of anesthesia and was released at the exact site of capture. A clean, baited trap was replaced at the site, and the original trap was returned to the processing site to be decontaminated before reuse. Animals from the control webs were treated similarly, except that blood and oral swab samples were not taken. Investigators recorded environmental data, including a general description of the vegetation and depending on resources available, more detailed descriptions of vegetation at individual webs and weather conditions during the trapping session (detailed rainfall and temperature data were available from meteorologic stations near each trapping site). Laboratory Analysis Serologic testing was conducted at CDC, Atlanta, or at the Arthropod-Borne and Infectious Diseases Laboratory, Colorado State University, Fort Collins, CO, USA. Samples of whole blood were tested for antibody reactive with SNV recombinant nucleocapsid protein antigen by enzyme-linked immunosorbent assay according to a standardized protocol (29). Briefly, blood specimens were initially diluted 1:25 in 5% skim milk in 0.01 M phosphate-buffered saline with 0.5% Tween-20 and subsequently diluted to 1:100 through 1:6,400 in fourfold dilutions in microtiter plates. Samples were tested against the recombinant nucleocapsid antigen and a recombinant control antigen (29). A conjugate mix of anti-Rattus norvegicus and anti-Peromyscus leucopus (heavy and light chains) immunoglobulin G (IgG) (Kirkegaard and Perry, Gaithersburg, MD) was used to detect bound immunoglobulin. Adjusted optical densities (OD) for each dilution were calculated by subtracting the OD(sup 410) of the control antigen from the OD(sup 410) of the SNV antigen. Titers were assigned on the basis of an adjusted OD value exceeding 0.20 for each dilution. A second measure consisting of the sum of the adjusted OD values for all four dilutions was also calculated. Serum specimens were considered SNV-positive if their titer was 1:400 or their sum-adjusted OD was 0.95. The cut-off values were determined by assessment of rodents found to be SNV-positive by several serologic tests during the initial investigation of the 1993 outbreak (13) and have been reassessed periodically among large populations of rodents collected in North and South America. Antibodies to other North American hantaviruses are cross-reactive with SNV antigen. This assay would detect (but not distinguish among) infections by New York virus (from the white-footed mouse [15]), Prospect Hill-like viruses (from arvicoline rodents [30]), El Moro Canyon virus (from the Western harvest mouse [31]), Black Creek Canal virus (from the cotton rat [16]), and Bayou virus (from the rice rat [17]). At this writing, analyses on oral swab specimens (antigen or antibody detection, polymerase chain reaction) have not been conducted. Blood and oral swab samples are archived in -70°C freezers at CDC, Atlanta, and at the Museum of Southwestern Biology, Albuquerque, New Mexico. Acknowledgments We are indebted to many investigators at CDC for laboratory analyses and database management. Special thanks to R. Meyer, M.L. Martin, V. Semenova, G. Gallucci, M. Curtis, A.J. Williams, and P. Stockton in the laboratory; B. Farrar in the stock rooms; and L. Morgan and K. Schmidt at the computer. K. Wagoner, K. Colbert, and B. Ellis prepared the figures. K. Abbott, C. Calisher, B. Ellis, and M. Morrison provided helpful comments on an earlier version of the manuscript. Dr. Mills is chief of the Medical Ecology Unit, Special Pathogens Branch, Division of Viral and Rickettsial Diseases, CDC. His research interests include zoonotic diseases, specifically host-pathogen evolution and interactions. Address for correspondence: James N. Mills, Centers for Disease Control and Prevention, 1600 Clifton Road, Mail Stop G14, Atlanta, GA 30333, USA; fax: 404-639-1118; e-mail: jum0@cdc.gov. References 1. Schmaljohn CS, Hjelle B. Hantaviruses: a global disease problem. Emerg Infect Dis 1997;3:95-104. 2. Peters CJ, Mills JN, Spiropoulou C, Zaki SR, Rollin PE. Hantaviruses. In: Guerrant RL, Walker DH, Weller PF, editors. Tropical infectious diseases: principles, pathogens, and practice. New York: W.B. Saunders. In press 1999. 3. Lee HW, French GR, Lee PW, Baek LJ, Tsuchiya K, Foulke RS. Observations on natural and laboratory infection of rodents with the etiologic agent of Korean hemorrhagic fever. Am J Trop Med Hyg 1981;30:477-82. 4. Hutchinson KL, Rollin PE, Peters CJ. Pathogenesis of a North American hantavirus, Black Creek Canal virus, in experimentally infected Sigmodon hispidus. Am J Trop Med Hyg 1998;59:58-65. 5. Tsai TF. 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Long-term studies of vertebrate communities. San Diego: Academic Press; 1996. p. 1-15. 21. Childs JE, Glass GE, Korch GW, LeDuc JW. Prospective seroepidemiology of hantaviruses and population dynamics of small mammal communities of Baltimore, Maryland. Am J Trop Med Hyg 1987;37:648-62. 22. Niklasson B, Hornfeldt B, Lundkvist A, Bjorsten S, LeDuc J. Temporal dynamics of Puumala virus antibody prevalence in voles and of nephropathia epidemica incidence in humans. Am J Trop Med Hyg 1995;53:134-40. 23. Mills JN, Ellis BA, McKee KT, Calderón GE, Maiztegui JI, Nelson GO, et al. A longitudinal study of Junín virus activity in the rodent reservoir of Argentine hemorrhagic fever. Am J Trop Med Hyg 1992;47:749-63. 24. Parmenter RR, Brunt JW, Moore DI, Ernest S. The hantavirus epidemic in the Southwest: rodent population dynamics and the implications for transmission of hantavirus-associated adult respiratory distress syndrome (HARDS) in the Four Corners region. Publication No.: 41. Albuquerque (NM): University of New Mexico; 1993. Sevilleta Long-Term Ecological Research Site. 25. Parmenter CA, Yates TL, Parmenter RR, Mills JN, Childs JE, Campbell ML, et al. Small mammal survival and trapability in mark-recapture monitoring programs for hantavirus. J Wildl Dis 1998;34:1-12. 26. Swann DE, Kuenzi AJ, Morrison ML, DeStefano S. Effects of sampling blood on survival of small mammals. Journal of Mammalogy 1997;78:908-13. 27. Mills JN, Yates TL, Childs JE, Parmenter RR, Ksiazek TG, Rollin PE, et al. Guidelines for working with rodents potentially infected with hantavirus. Journal of Mammalogy 1995;76:716-22. 28. Mills JN, Childs JE, Ksiazek TG, Peters CJ, Velleca WM. Methods for trapping and sampling small mammals for virologic testing. Atlanta (GA): U.S. Department of Health and Human Services; 1995. 29. Feldmann H, Sanchez A, Morzunov S, Spiropoulou CF, Rollin PE, Ksiazek TG, et al. Utilization of autopsy RNA for the synthesis of the nucleocapsid antigen of a newly recognized virus associated with hantavirus pulmonary syndrome. Virus Res 1993;30:351-67. 30. Yanagihara R, Daum CA, Lee PW, Baek LJ, Amyx HL, Gajdusek DC, et al. Serological survey of Prospect Hill virus infection in indigenous wild rodents in the USA. Trans R Soc Trop Med Hyg 1987;81:42-5. 31. Hjelle B, Chavez-Giles F, Torrez-Martinez N, Yates T, Sarisky J, Webb J, et al. Genetic identification of a novel hantavirus of the harvest mouse Reithrodontomys megalotis. J Virol 1994;68:6751-4. —————————————————————————————————————————————————————————————————————————— Hantavirus Long-Term Hantavirus Persistence in Rodent Populations in Central Arizona Ken D. Abbott,* Thomas G. Ksiazek,† and James N. Mills† *Yavapai College, Prescott, Arizona, USA; and †Centers for Disease Control and Prevention, Atlanta, Georgia, USA --------------------------------------------------------------------------- For 35 months, we monitored hantavirus activity in rodent populations in central Arizona. The most frequently captured hantavirus antibody–positive rodents were Peromyscus boylii and P. truei. Antibody-positive P. boylii were more frequently male (84%), older, and heavier, and they survived longer on trapping web sites than antibody-negative mice. The number of antibody-positive P. boylii was greater during high population densities than during low densities, while antibody prevalence was greater during low population densities. Virus transmission and incidence rates, also related to population densities, varied by trapping site. The spatial distribution of antibody-positive P. boylii varied by population density and reflected the species preference for dense chaparral habitats. The focal ranges of antibody-positive P. boylii also demonstrated a patchy distribution of hantavirus. We report results of the initial 35 months of one of several longitudinal hantavirus studies begun in the southwestern United States after the 1993 outbreak of hantavirus pulmonary syndrome (HPS) (Mills et al., this issue, pp. 95-101). This study monitors and quantifies the seasonal and year-to-year changes in rodent populations and the prevalence and incidence of hantavirus infection, identifies environmental factors associated with these dynamics, explores aspects of temporal and spatial viral transmission within reservoir populations, and examines the characteristics of infected animals. Trapping and Processing In January 1995, we established four 3.14-ha mark-recapture trapping webs in northcentral Arizona, elevation 1,648 m (Mills et al., this issue, pp. 95-101). The webs were located north of Prescott in Limestone Canyon (35°31'N, 121°29'W). All sites were in juniper-pinyon and interior chaparral communities (1), although each site varied in physiognomy, aspect, slope, and plant species composition and distribution. Trapping web sites S-1 and C-1 were separated by a valley 150 m wide and were .6 km north of sites S-2 and C-2, which were set apart by a 100-m ravine and creek bed. All webs were operated from January 1995 to September 1996. Serologic samples were taken from rodents captured at S-1 and S-2, while C-1 and C-2 were initially operated as control sites to determine the effects of sampling on rodent survivorship. In October 1996, trapping was discontinued at C-2 (since our field data and others' [2] indicated that sampling had no effect on rodent survival), and blood collection and antibody testing were initiated at C-1 because of its microhabitat uniqueness and high rodent densities. Web design and placement, trapping periods, mark-recapture techniques, animal processing, and serologic sampling procedures are described in Mills et al. (this issue, pp. 95-101). We anesthetized animals by securing the dorsal skin behind the head and slipping a nose cone with cotton wetted with isoflorane over the nose. Between animals, the nose cone was cleaned with disinfectant. When clearly anesthetized, the animal was placed on a clean table, measured, ear-tagged, and bled. Serologic testing was conducted at the Centers for Disease Control and Prevention, Atlanta, Georgia. Samples of whole blood were tested for antibody reactive with Sin Nombre virus (SNV)-recombinant nucleocapsid protein antigen by enzyme-linked immunosorbent assay (ELISA) (3). The laboratory methods we used are described in Mills et al.; (this issue, pp. 95-101). Data Analysis Peromyscus boylii (brush mouse) and P. truei (pinyon mouse) were assigned to three categories on the basis of body mass at first capture. Body mass classes (derived from our field data and other sources [4]) were used as an indication of relative age: 6.0 g to 19.0 g (juvenile), 19.1 g to 22.0 g (young adult), and 22.1 g to >30.0 g (adult). We estimated the survival of trappable populations by using mark-recapture data to assess the number of times an animal was caught between the first and last capture. While not a measure of actual life span, average survival provides some indication of population turnover and longevity (5). The minimum number alive (the number of rodents captured in a month plus the number of rodents captured on at least one prior and one subsequent occasion) was used to estimate population sizes (5-7). The minimum number infected was calculated for antibody-positive rodents by using the same technique. Estimated standing prevalence was calculated by dividing the monthly minimum number infected by minimum number alive. These methods provide an estimate of the number of rodents alive and population sizes for a period, an estimate of the number of infected rodents, and comparisons of antibody prevalence between trapping web locations. Field data were transferred to a computer database by using Excel (Microsoft Corp., Redmond, WA) and Lotus 1-2-3 for Macintosh (Lotus Development Corporation, Cambridge, MA). Statistical analyses were performed by using MINITAB (Minitab Inc, State College, PA) statistical software, the Mann-Whitney and two-sample t tests, one-way analysis of variance, and linear trend model (8). Trapping Results During 35 months of trapping at three grids, 844 rodents were captured 3,552 times. Blood samples were obtained from 553; from these rodents, 1,418 samples were collected (as a result of subsequent captures of the same rodents during progressive trapping sessions) and tested for hantavirus antibody (Table 1). Table 1. Sin Nombre virus–antibody-positive mice and hantavirus prevalence at three mark-recapture webs, December 1995–November 1997(sup a) -------------------------------------------------------------------------- Trapping webs ----------------------------------- Species S-1 S-2 C-1(sup b) Totals -------------------------------------------------------------------------- Peromyscus boylii 76/286/109 74/516/178 3/56/22 153/858/309 (Brush mouse) (26.6%) (14.3%) (5.4%) (17.8%) Peromyscus truei 3/165/67 5/133/55 0/15/8 8/313/130 (Pinyon mouse) (2.0%) (3.8%) (0.0%) (2.6%) Tamias dorsalis 0/73/40 0/83/29 0/19/9 0/175/78 (Cliff chipmunk) (0.0%) Dipodomys ordii 0/3/2 0/33/13 0/7/3 0/43/18 (Ord's kangaroo rat) (0.0%) Onychomys leucogaster 0 0/10/3 0 0/10/3 (Northern grasshopper (0.0%) mouse) Neotoma stephensi 0/3/1 0/2/2 0/3/2 0/8/5 (Stephen's woodrat) (0.0%) Neotoma albigula 0/1/4 0/4/1 0/2/1 0/7/6 (White-throated (0.0%) wood rat) Reithrodontomys megalotis 0 0/4/4 0 0/4/4 (Western harvest mouse) (0.0%) All species 79/531/223 79/785/285 3/102/45 161/1,418/553 -------------------------------------------------------------------------- (sup a)Positive samples/number of samples tested/number individuals tested. Values in parentheses are hantavirus antibody prevalences for 35 months based on the number of samples tested. (sup b)C-1 was initially a control web; serologic sampling began in October 1996. P. boylii was the most commonly captured species (70%), followed by P. truei (18%), Tamias dorsalis (9%), and Dipodomys ordii (2%). Irregular species (Neotoma albigula, N. stephensi, Onychomys leucogaster, and Reithrodontomys megalotis) accounted for 1% of the total captures. The highest rodent densities occurred at webs S-2 and C-1 (40% and 33% of all captures, respectively), while S-1 accounted for 27% of the total captures. Population Dynamics Population levels of the two most frequently captured rodent species, P. boylii and P. truei, were relatively high through the winter of 1995 to 1996 and then declined (p <0.05) during the subsequent summer and autumn, remaining at low levels through 1996 to 1997 (Figure 1). The P. boylii population had the most persistent decline (76%) followed by T. dorsalis (64%) and P. truei (34% short-term reduction). Population levels of P. boylii were consistently higher than those of P. truei, except for the summer of 1997 (May through August); during this period P. boylii densities were at their lowest, 6.5 animals per 6.2 ha per month, while the P. truei populations increased to near high density levels (12.2 animals per 6.2 ha per month). For 4 months, far more P. truei were captured than P. boylii (Figure 1). [Fig] Figure 1. Minimum number of Peromyscus boylii and P. truei alive (MNA) and the minimum number infected (MNI) with Sin Nombre virus (antibody-positive) at two mark-recapture webs (6.2 ha).* *Because of adverse weather conditions, we only trapped for 2 nights in January and May 1995. During the first 5 months, adverse weather conditions (rain, snow, high winds) hampered trapping efforts. Strong wind and wind gusts seemed the main factor contributing to reduced periodic capture rates (Figure 1). Characteristics of Antibody-Positive Captured Rodents Although data from C-1 were not included in comparative analysis because serologic sampling was not initiated at this site until October 1996 during low population densities (4.0 samples per month, range 0 to 8), of the 21 P. boylii captured and tested, 2 were hantavirus–antibody- positive (10%); 0 (0%) of 7 females and 2 (28%) of 14 males. After samples were collected from one antibody-positive P. boylii in October 1996, no antibody-positive samples were collected until the following October, when another P. boylii, which had survived for 12 months, became antibody- positive for the first time. The 62 hantavirus antibody–positive rodents captured at the two sites represented two species: 58 P. boylii and 4 P. truei (Table 2). The prevalence of hantavirus antibody differed considerably by species: P. boylii had a prevalence of 20%, P. truei 3%. All four antibody-positive P. truei were trapped before September 1996 when population densities were high for all rodent species. Table 2. Antibody-positive and antibody-negative Peromyscus boylii and P. truei at two mark-recapture webs,(sup a) December 1995—November 1997 -------------------------------------------------------------------------- No. (%) P. boyii No. (%) P. truei --------------------------- ---------------------------- Characteristic Positive Negative Totals Positive Negative Totals -------------------------------------------------------------------------- Sex Male 49 (32) 106 (68) 155 (54) 3 (4) 63 (96) 66 (56) Female 9 (7) 123 (93) 132 (46) 1 (2) 50 (98) 51 (44) Totals 58 (20) 229 (80) 287 4 (3) 113 (97) 117 Body mass class(sup b) I 2 (3) 75 (97) 77 (27) 0 31 (100) 31 (26) II 9 (15) 51 (85) 60 (21) 1 (5) 18 (95) 19 (16) III 47 (31) 103 (69) 150 (52) 3 (4) 64 (96) 67 (58) Web-site longevity [months](sup c) Male 4.4[1-16] 2.9[1-26] 2.3[1-5] 3.2[1-18] Female 3.3[1-13] 3.5[1-18] 1 [1] 3 [1-15] -------------------------------------------------------------------------- (sup a)S-1 and S-2 webs. (sup b)Classes assigned at first capture. I = 6.0g-19.0g; II = 19.1g-22.0g; III = 22.1g to >30.0g. (sup c)Longevity is the mean number of months animals were captured, from first to last capture. Values in brackets are ranges. Antibody-positive Peromyscus were more often male and within the heaviest mass class (Table 2). Although approximately half of the P. boylii tested were male, 84% of the antibody-positive mice were male. The male-to-female ratio was similar to that of P. truei, despite the small sample size. We found more adults and fewer young among the antibody-positive Peromyscus, even though young-to-adult capture ratios were similar among seronegative mice. Longevity of antibody-positive mice was considerably different between the two species, while longevity of antibody-negative mice was similar (Table 2). Antibody-positive male P. boylii tended to survive longer than antibody-positive female. Furthermore, antibody-positive male P. boylii lived longer than antibody-negative male P. boylii (4.4 months and 2.9 months, respectively; t = 2.58, df = 48, p = 0.007). P. boylii Population Dynamics and Temporal Patterns of Infection The number of captures per month and the number of samples per month were usually not the same—some animals were not sampled because of death, weakened physical condition, hypothermia, or escape. The number of animals tested for antibody to hantavirus, however, mirrored population trends. The P. boylii population declined dramatically during summer and autumn 1996, stabilized at low levels during winter 1996 and 1997, and fell to minimal levels in spring 1997 (Figure 1) (Table 3). Table 3. Population densities and hantavirus-antibody prevalence in Peromyscus boylii at two mark-recapture trapping webs, by period ------------------------------------------------------------------------------ Dec 1995-Nov 1997 High density(sup a) Low density(sup b) ---------------------- --------------------- ------------------------ Density/ Prevalence/ Density/ Prevalence/ Mean/ Prevalence/ Web month month month month month month sites (sup c) (sup d) (sup c) (sup d) (sup c) (sup d) ------------------------------------------------------------------------------ S-1 & 26.1 20.2 43.6 18.4 11 25.4 S-2 (4-52) (0-43) (32-52) (10-22) (4-22) (12-43) S-1 9.7 28.5 15.2 26.4 4.5 37.0 (1-20) (0-75) (10-20) (15-38) (1-9) (0-75) S-2 16.4 14.2 28.4 14.3 6.5 15.0 (3-42) (0-33) (13-42) (6-19) (3-13) (0-33) ------------------------------------------------------------------------------ (sup a)June 1995 to June 1996 (sup b)September 1996 to September 1997 (sup c)Population density (number of individuals per 6.2 hectares), determined by minimum number alive. Values in parentheses are ranges. (sup d)Antibody prevalence to hantavirus (%), determined by estimated standing prevalence. Values in parentheses are ranges. [Fig] Figure 2. Minimum number of living Peromyscus boylii and the estimated standing prevalence of hantavirus antibody–positive mice at two mark-recapture webs (6.2 ha).* *Because of adverse weather conditions, we only trapped for 2 nights in January and May 1995. For the 35-month sampling period, the mean number of antibody-positive P. boylii was 5.0 animals per 6.2 ha per month, range 0 to 11 (Figure 1). The number of antibody-positive P. boylii was higher during high population densities than during low densities (8.0 and 2.8 animals per 6.2 ha per month, respectively; t = 4.83, df = 21, p < 0.001). Numbers of antibody-positive animals were similar during 35 months at S-1 and S-2 (2.7 and 2.4 animals per 6.2 ha per month, respectively), even though population densities at S-2 were regularly higher than at S-1. The mean antibody prevalence for the sampling period was 20.2% (range 0% to 43%) and was higher during low densities than high densities (Figure 2). At each site, antibody prevalence rates were also higher during low densities, but not significantly different from rates during high population densities. However, antibody prevalence varied between sites and was consistently higher at S-1 (Table 3). The highest mean monthly antibody prevalence occurred on S-1 during low population densities (37.0%) and was higher than prevalence on S-2 during the same period. The highest monthly antibody prevalence occurred at S-1 during minimal population densities, May 1997, when three of four captured P. boylii were antibody-positive (75%). Maximum and minimum antibody prevalence occurred during similar periods at both sites. During low population densities, S-2 had 4 months without an antibody-positive sample, while antibody-positive animals were not captured from S-1 for 2 months. The months when no antibody-positive animals were captured were not the same for both sites; at least one positive sample was recorded each month, even during low population densities. Longevity and Seroconversion of Infected Mice Hantavirus antibody-positive P. boylii tended to survive longer (mean 4.2 months) than seronegative mice (mean 3.2 months) (t = 1.77, df = 138, p = 0.04) (Table 4). At site S-2, survival was similar between antibody-positive and antibody-negative mice, but at S-1, antibody-positive mice lived longer (4.8 months) than seronegative mice (3.0 months) (t = 2.58, df = 48, p = 0.007). At both sites, survival among male and female mice was not significantly different. Table 4. Frequency of intervals between first and last capture of individual Peromyscus boylii, December 1995–November 1997 ------------------------------------------------------------------------------------------ No. months in interval between first and last captures -------------------------------------------------------- No. Web sites P.boylii 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 18 19 26 Mean (sup a) (sup b) ------------------------------------------------------------------------------------------ Antibody-positive mice S-1 & S-2 58 22 4 6 5 7 2 2 2 3 2 1 1 1 4.2 S-1 30 10 2 2 1 6 1 1 2 2 1 1 1 4.8 S-2 28 12 2 4 4 1 1 1 1 1 1 3.5 Antibody negative mice S-1& S-2 250 117 48 24 11 11 6 7 7 3 4 3 3 1 2 1 1 1 3.2 S-1 90 47 15 8 3 4 2 2 2 2 3 1 1 3.0 S-2 160 70 33 16 8 7 4 5 5 1 4 2 1 2 1 1 3.3 ------------------------------------------------------------------------------------------ (sup a)Total number of individual P. boylii. (sup b)Mean number of months in interval. [Fig] Figure 3. Initial antibody acquisition in Peromyscus boylii at two mark- recapture webs, by month, December 1995–November 1997. Initial acquisition of hantavirus antibody (seroconversion) was observed in 33% of the antibody-positive P. boylii. P. boylii acquired hantavirus antibody in all months except December, January, and March (Figure 3). Two transmission peaks, accounting for 79% of seroconversions, took place during the typical 7-month reproductive period, April through October (37% during April, May, and June; 42% during September and October). Seroconversions at S-2 were directly related to population levels, with 9 (90%) of 10 S-2 seroconversions taking place during high population densities in 1995. This relationship did not appear at S-1, where the number of seroconversions was similar during high and low population densities. Incidence of Infection The incidence rate for seroconversion per 100 mice per month was twice as high at S-2 (34.5) as at S-1 (17.0). The greater number of mice at risk and the number of months before seroconversion accounted for the higher incidence rate at S-2 (Table 5). S-1 had fewer P. boylii at risk, which did not seroconvert for an average of 5.4 months; S-2 had a larger number of mice at risk, which seroconverted after 2.2 months. Table 5. Incidence rates of hantavirus infection in Peromyscus boylii that were recaptured and sampled at least twice, December 1995–November 1997, two web sites ------------------------------------------------------------------------ No. Mouse-mo. Mean mo. at (Cumu of obser- before risk Serocon- lative vation Incidence serocon- Sites (sup a) versions %) (sup b) (sup c) version ------------------------------------------------------------------------ S-1 43 9 (20.9) 53.0 17.0 5.4 S-2 90 10 (11.1) 29.0 34.5 2.2 S-1 & S-2 133 19 (14.3) 82.0 23.2 3.7 ------------------------------------------------------------------------ (sup a)Antibody-negative at time of first capture. (sup b)Includes all time intervals between successive captures when mice were seronegative, and half the interval between captures when mice became seropositive. (sup c)Seroconversions per 100 mice per month. Spatial of Patterns of Infected Mice Distribution and movement of antibody-negative and antibody-positive P. boylii varied by population density and availability of sheler and food resources. At both trapping web sites, P. boylii distributions were associated with brushy chaparral plant species. The ranges of high density mice outline, in general, the distribution of thick chaparral stands (Figure 4). Because plant species diversity and belts of chaparral stands were greater at S-2, P. boylii distribution was relatively continuous and widespread. Chaparral stands at S-1 were discontinuous, and P. boylii lived in rocky pockets of vegetation and were seldom trapped in different chaparral pockets if separated by open terrain (Figure 4). At both sites, P. boylii avoided open juniper-pinyon areas. [Fig] Figure 4. Ranges and trap stations of hantavirus antibody–positive Peromyscus boylii during high and low population densities. Each web covered 3.1 ha. Trap stations within ranges were occupied by antibody-negative and antibody-positive mice at various times. High densities represent 13 months (June 1995 to June 1996), and low densities represent 13 months (September 1996 to September 1997). During periods of high population density, antibody-positive mice occupied scattered chaparral habitats of undergrowth areas of the sites and moved freely between web transects. Movement, however, appeared to be directly influenced by chaparral cover. During low population densities, antibody-positive mice withdrew to a few, well-defined refuges (Figure 4). The movement of antibody-positive P. boylii during low densities was also restricted; mice seldom moved between web transects. Hantavirus Prevalence Rates and Patterns The prevalence rates of P. boylii (20.2%) and P. truei (3%) in our study were similar to those found in other studies carried out in pinyon-juniper habitats (4). The short-term infection in P. truei may have been caused by spillover from syntopic P. boylii (the four antibody-positive P. truei were found only during spring and summer 1995, when P. boylii densities and the potential of interspecies contact were greatest). Six other rodent species coexisting with P. boylii should have had similar risks for hantaviral infection since they had been captured at trap stations used by P. boylii at one time or another (capturing two or three different species at one station during a single trapping session was not uncommon). The evident rarity of hantavirus infection in P. truei and the absence of infection in other sympatric rodents suggests that P. boylii is the primary hantavirus host in this area and that transmission to other rodent species may be unlikely during periods of average population densities. Similar relationships have been demonstrated in southern Arizona, where only species of Peromyscus had immunoglobulin G (IgG) antibody reactive with SNV (Kuenzi et al., this issue, pp. 113-117). On the basis of long-term infection patterns and persistent virus shedding (9-11), we assume that hantavirus antibody–positive P. boylii are chronically infected and infectious (Mills et al., this issue, pp. 135-142). Studies using reverse transcription-polymerase chain reaction (RT-PCR) on blood samples from field-caught P. maniculatus from Nevada (12) mirror other studies of host-hantavirus associations in suggesting initial viremia, followed by a relatively rapid immune response that cleared virus from blood in approximately 1 month (animals remained antibody-positive for at least 7 months). However, as numerous studies have shown (9-11;13), the short duration of hantavirus RNA in blood does not reflect its residence in organs. Another study demonstrated that 97% of antibody-positive P. maniculatus were PCR-positive for viral RNA in organ tissues (13), which implies chronic infection, as has been demonstrated for other hantavirus-host associations. Nevertheless, the crucial experiments to demonstrate chronic infection and persistent shedding have not been done for P. boylii. We are attempting to develop methods to reliably and consistently collect urine from mark-release-capture animals in the field to address this problem. Slightly more male than female mice (1.2:1) were tested for antibody to hantavirus; however, fewer male than female mice (1:1.2) were antibody-negative. The higher antibody prevalence in males may be due to territoriality, aggression toward other males during breeding periods, longer survival, and breadth of travel (4,14). Factors Affecting Population Density The population densities and distributions of rodents were related to seasonal and year-to-year availability of acorns, seeds, and juniper berries (mast). Acorns, pinyon seeds, juniper berries, and grasses were abundant throughout the study sites during summer and autumn 1995, reflecting surplus winter precipitation the previous 3 years (15). Population levels of the rodent community—relatively high during autumn and winter 1995-96—may have been related to this abundance of seed crops. During the winters of 1995-96 and 1996-97, precipitation was well below normal, and the first winter drought resulted in complete mast failures by all chaparral species and pinyon pine. Juniper crops, evident in autumn 1996, were depleted soon after. The second winter drought resulted in the mast failure of oak species, pinyon, and juniper, but other chaparral species produced minimal crops during the spring and late summer (Abbott et al., unpub. data). P. boylii population fluctuations (Figure 1), related to year-to-year mast resources and variations in seasonal female reproductive efforts, are consistent with fluctuations of mast-consuming Peromyscus and Tamias rodents, which show a positive correlation between mast production and breeding behavior (16-18). Comparable regional population fluctuations occurred during this same period in Colorado and southern Arizona (Calisher et al., this issue, pp. 126-134; Kuenzi et al., this issue, pp. 113-117). Female reproductive activity was consistently absent during the colder winter months of November through February. The reproductive period, April through October, typically unimodal and coinciding with seed development of syntopic vegetation, started at low levels in April and peaked in late summer and autumn. Mast of oak species, pinyon pine, and juniper usually ripen in late summer and early autumn while summer monsoons may cause other chaparral species to produce seeds in both spring and summer (Abbott et al., unpub. data). This pattern of reproduction and food supply was evident during the 1995 breeding season; 40% of the female mice captured in spring showed signs of reproductive activity, compared with 76% of those captured in summer and 86% of those captured in autumn. The subsequent breeding season began normally, with 43% of females pregnant, but new pregnancies nearly halted during the summer months, decreasing 96% from the previous year. Only a few of the 64 female mice captured in June and July 1996 had a perforate vagina, and none showed signs of lactation or pregnancy. The autumn breeding effort declined by 56% from the previous year. During the 1997 breeding season, there were 84% fewer female mice than during the 2 previous years, but most were reproductively active, suggesting that a population recovery was under way. Factors Affecting Hantavirus Prevalence The number of hantavirus-infected mice was higher during the high population densities of 1995-96 (Figure 1). Month-to-month numbers of antibody-positive mice appeared more stable than those of antibody-negative mice. The number of high density–antibody-positive P. boylii was stable during the winter, with small peaks proportional to monthly capture success. The numbers of antibody-positive mice remained stable (though lower) during the subsequent precipitous 7-month population decline. Even during low population densities, antibody-positive mice were persistent at minimal, yet stable levels. This consistent presence of at least a few infected mice may reflect the resident nature of antibody-positive mice, characteristically older and able to survive for longer periods. Fifty-five percent of the antibody-positive mice survived on trapping web sites 3 months or longer and were considered resident, while 34% of the seronegative mice were resident. The proportion of hantavirus antibody–positive P. boylii varied by population density and trapping web site (Figure 2) (Table 3). S-1 maintained the highest mean antibody prevalence; during low population densities, prevalence increased. Almost half of the P. boylii captured at S-1 tested positive during low density months when at least one mouse was antibody-positive. Population densities at S-2 were consistently greater than at S-1 and were associated with lower overall prevalence rates. Approximately 23% of the P. boylii captured at S-2 were antibody-positive during low density months when at least one mouse was antibody-positive. Positive linear correlations between population density and antibody prevalence have not been found in other species of Peromyscus (Calisher et al., this issue; pp. 126-134;11;19). We observed that one third of the antibody-positive P. boylii acquired antibody. No mice reverted from antibody-positive to antibody-negative. Transmission of hantavirus was bimodal and associated with spring and autumn reproductive activity (Figure 3). Thirty-seven percent of P. boylii seroconverted in the spring, and 42% in the autumn reproductive period. Mice that seroconverted were more frequently male, within the heaviest mass class, and survived longer than mice that remained antibody-negative. The trend for bimodal transmission may reflect intraspecific competition, greater movement, and aggressive behavior by resident antibody-positive males during peak reproductive periods (20). Similar transmission trends have been reported in rat populations (6). Consequently, risks of horizontal transmission may increase during the more active seasons. Incidence of infection varied with population densities, recapture rates, and population dynamics. Rates of P. boylii seroconversion varied by site, but collectively, both sites had an average 14.3% incidence of infection among the population at risk during the study period (Table 5). The number of seroconversions at both sites was similar, but the number of mice at risk at S-2 was much larger, since population densities were regularly higher. Consequently, the cumulative proportion of mice seroconverting at S-2 was 47% lower than at S-1, whereas the incidence of seroconversions per 100 mice per month was 103% greater. Characteristics of the S-1 population (longer survival as antibody-negative animals, more restricted centers of activity, and continuous infection during periods of high and low population densities) may have been contributing factors to the difference in incidence rates between sites. The focal ranges of antibody-positive P. boylii were patchy; they expanded and contracted over time (Figure 4). Hantavirus infection and distribution patterns were influenced by habitat structure, seasonal food availability, and the behavioral characteristics of infected mice. At both sites, P. boylii were associated with corridors and patches of chaparral understory within the juniper-pinyon woodland, and especially with dense stands of chaparral associated with rocky substrates and downed trees that provided optimal shelter. These favored sites were usually located on slopes and along creek channels. In southern Arizona, P. boylii were found in analogous habitat distributions; the species favored oak riparian vegetation, and most were captured in one portion of one trapping web (Kuenzi et al., this issue, pp. 113-117). Diverse chaparral stands were more widespread and continuous at S-2. During high population densities, P. boylii occupied scattered chaparral areas throughout most of the web and were often trapped at sites several meters apart. The relatively high abundance of mice over a large area may explain the greater incidence of infection and lower antibody prevalence at S-2. The greater number of mice during high population densities and the greater turnover rate seemed to dilute the prevalence of infection and, at the same time, increase the risk for infection because of intensified encounters. The patchiness of hantavirus infection was more evident and focalized at S-1. Chaparral stands were discontinuous; P. boylii occupied discrete chaparral pockets, seldom migrating from one pocket to another (Figure 4). During this study, S-1 had three prominent centers of hantavirus infection and three associated centers of P. boylii activity. The structure and disjunct nature of the activity centers (and associated centers of antibody-positive animals) may have contributed to higher antibody prevalences and greater cumulative seroconversion since the mice occupying these restricted habitats had a greater chance of encountering each other. During low population densities, the higher prevalence rates of 50% to 75% were related to antibody-positive male mice that were older, heavier, and able to reside for a longer period within the activity centers. Similar patterns of clustering or patchiness and hantavirus infection have been documented for cotton rats, Sigmodon hispidus, in Florida (21). Along with high population densities, the longer stay of dominant male mice in optimal and reliable habitats may be a primary variable contributing to hantavirus infection. This assumption is based on three trends: animals that became antibody-positive survived longer than those that did not seroconvert; antibody-positive tended to survive longer than antibody-negative mice; and in patchy optimal habitats, resident mice tended to be dominant, male, and antibody-positive. Consequently, resident male mice may provide a reliable reservoir during low population densities and therefore ensure the survival of hantavirus within rodent communities. Conclusions Our preliminary results, and those of other recent studies (Kuenzi et al.,this issue, pp. 113-117;18), have implicated precipitation, habitat structure, and food resources as ultimate environmental factors that influence reservoir population dynamics, viral transmission, and hantavirus persistence. The results of this and other recent studies have raised questions concerning proximate patterns of hantavirus maintenance, seroconversions, and transmission within specific reservoir species occupying different western regions (Mills et al., this issue, pp. 135-142). Additional data suggesting that sex ratios, size, and social organization affect temporal and spatial seroconversion relationships will be addressed in forthcoming articles. We hope that this ongoing study will collect sufficient data to explain the interplay of habitat resources, social hierarchies, intraspecific competition, and dispersal behavior and how these proximate factors influence hantavirus ecology and human risk. Acknowledgments Special recognition is due to field crew regulars who endured extreme field conditions: Lisa Gelczis, Nathan Zorich, Tyler Williams, Samantha Yazzi, Henry Provencio, Dan Carroll, Heather Shane, Chris Davis, Jon Mock, Karen Mock, and Romey Haberle. Thanks to E. Chambers for veterinary guidance, C. Levy for initial assistance, B. Farrar for shipping supplies when requested, J. Dunnum and C. Parmenter for sample management, C.J. Peters, J.E. Childs, and two anonymous reviewers for manuscript suggestions, D. Dailey for valuable support throughout this study, and the Chino Ranger District, Prescott National Forest, for necessary permits. This work was supported by grants 08-5071 from the Arizona Department of Health Services and U50/CCU913429-02 from the Department of Health and Human Services, Public Health Service, Centers for Disease Control and Prevention. Partial support was provided by the Yavapai College Foundation. Dr. Abbott is chair of the Biology Department, Yavapai College, Prescott, Arizona. His research focuses on small mammal population ecology and hantavirus associations, vertebrate metapopulation distributions and ecology, and riparian ecology. His areas of expertise include desert ecology and the physiologic ecology of vertebrates; he serves as an ecologic consultant to federal, state, and private agencies. Address for correspondence: Ken D. Abbott, Department of Biology, Yavapai College, 1100 E. Sheldon, Prescott, AZ 86301, USA; fax: 520-776-2315; e-mail: sm_ken@yavapai.cc.az.us. References 1. Brown DE, editor. Biotic communities. Southwestern United States and Northwestern Mexico. Salt Lake City: University of Utah Press; 1994. 2. Swann DE, Kuenzi AJ, Morrison ML, DeStefano S. Effects of sampling blood on survival of small mammals. Journal of Mammalogy 1997;78:908-13. 3. Feldman H, Sanchez A, Morzunov S, Spiropoulou CF, Rollin PE, Ksiazek TG, et al. Utilization of autopsy RNA for the synthesis of the nucleocapsid antigen of a newly recognized virus associated with hantavirus pulmonary syndrome. Virus Res 1993;30:351-67. 4. Mills JN, Ksiazek TG, Ellis BA, Rollin PE, Nichol ST, Yates TL, et al. Patterns of association with host and habitat: antibody reactive with Sin Nombre virus in small mammals in the major biotic communities of the southwestern United States. Am J Trop Med Hyg 1997;56:272-84. 5. Mares MA, Ernest KA. Population and community ecology of small mammals in a gallery forest of central Brazil. Journal of Mammalogy 1995:76:750-68. 6. Childs JE, Glass GE, Korach GW, LeDuc JW. Prospective seroepidemiology of hantaviruses and population dynamics of small mammal communities from Baltimore, Maryland. Am J Trop Med Hyg 1987;37:648-62. 7. Krebs CJ. Demographic changes in fluctuating populations of Microtus californicus. Ecological Monographs 1966;36:239-73. 8. Zar JH. Biostatistical analysis. 3rd ed. Englewood Cliffs: Prentice Hall, Inc.; 1996. 9. Lee HW, Lee PW, Baek LJ, Song CK, Seong IW. Intraspecific transmission of Hantaan virus, etiologic agent of Korean hemorrhagic fever, in the rodent Apodemus agrarius. Am J Trop Med Hyg 1981;30:1106-12. 10. Yanagihara R, Amyx HL, Gajdusek DC. Experimental infection with Puumala virus, the etiologic agent of nephropathia epidemica, in bank voles (Clethrionomys glareolus). J Virol 1985;55:34-8. 11. Hutchinson KL, Rollin PE, Peters CJ. Pathogenesis of a North American hantavirus, Black Creek Canal virus, in experimentally infected Sigmodon hispidus. Am J Trop Med Hyg 1998;59:58-65. 12. Boone JD, Otteson EW, McGwire KC, Villard P, Rowe JE, St Jeor SC. Ecology and demographics of hantavirus infections in rodent populations in the Walker River Basin of Nevada and California. Am J Trop Med Hyg 1998;59:445-51. 13. Childs JE, Ksiazek TG, Spiropoulou CF, Krebs JW, Morzunov S, Maupin GO, et al. Serologic and genetic identification of Peromyscus maniculatus as the primary rodent reservoir for a new hantavirus in the southwestern United States. J Infect Dis 1994;169:1271-80. 14. Mills JN, Ellis BA, McKee KT, Calderon GE, Maiztegui JI, Nelson GO, et al. A longitudinal study of Junin virus activity in the rodent reservoir of Argentine hemorrhagic fever. Am J Trop Med Hyg 1992;47:749-63. 15. Abbott K. The effects of drought and mast failure on rodent populations and Sin Nombre virus in central Arizona. Proceedings of the 4th International Conference on HFRS and Hantavirus; 1998 Mar 5-7; Atlanta, GA. p. 58. 16. Jameson EW. Reproduction of deer mice (Peromyscus maniculatus and P. boylii) in the Sierra Nevada, California. Journal of Mammalogy 1953;34:44-58. 17. Wolf JO. Population fluctuations of mast-eating rodents are correlated with production of acorns. Journal of Mammalogy 1996;77:850-6. 18. Gashwiler JS. Deer mouse reproduction and its relation to the seed crop. American Midland Naturalist 1979;102:95-104. 19. Douglas RJ, Van Horn R, Coffin K, Zanto SN. Hantavirus in Montana deer mouse populations: preliminary results. J Wildl Dis 1996;32527-30. 20. Vessey SH. Long-term population trends in white-footed mice and the impact of supplemental food and shelter. American Zoologist 1987;27:879-90. 21. Glass GE, Livingstone W, Mills JN, Illacly G, Fine JB, Biggler W, et al. Black Creek Canal virus infection in Sigmodon hispidus in southern Florida. Am J Trop Med Hyg. In press 1999. —————————————————————————————————————————————————————————————————————————— Hantavirus A Longitudinal Study of Sin Nombre Virus Prevalence in Rodents, Southeastern Arizona Amy J. Kuenzi, Michael L. Morrison, Don E. Swann, Paul C. Hardy, and Giselle T. Downard University of Arizona, Tucson, Arizona, USA --------------------------------------------------------------------------- We determined the prevalence of Sin Nombre virus antibodies in small mammals in southeastern Arizona. Of 1,234 rodents (from 13 species) captured each month from May through December 1995, only mice in the genus Peromyscus were seropositive. Antibody prevalence was 14.3% in 21 white-footed mice (P. leucopus), 13.3% in 98 brush mice (P. boylii), 0.8% in 118 cactus mice (P. eremicus), and 0% in 2 deer mice (P. maniculatus). Most antibody-positive mice were adult male Peromyscus captured close to one another early in the study. Population dynamics of brush mice suggest a correlation between population size and hantavirus-antibody prevalence. We examined the role of rodent species as natural reservoirs for hantaviruses in southeastern Arizona to identify the species infected with hantavirus, describe the characteristics of infected animals, and assess temporal and intraspecific variation in infection rates. Trapping Procedures Beginning in May 1995, we established four permanent trapping webs on the Santa Rita Experimental Range in the Santa Rita Mountains of southeastern Arizona (Pima County). The design of these webs, as well as details on mark-recapture trapping procedures, are described by Mills et al. (this issue, pp. 95-101). Elevations of the trapping webs are approximately 1,250 m to 1,379 m. All trapping webs contained approximately equal amounts of two main vegetation types, semidesert grassland (characterized by Lehmann lovegrass [Eragrostis lehmanniana], three-awn [Aristida spp.], prickly pear cactus [Opuntia spp.], and mesquite [Prosopis velutina]) and oak riparian (characterized by deciduous trees including Arizona white oak [Quercus arizonica] and netleaf hackberry [Celtis reticulata]; occurs in drainage areas where water flow is seasonally intermittent), occur at these elevations. Web 1 was operated from May 1995 through September 1996, when trapping was discontinued because of low trap success, and webs 2, 3, and 4 were operated from May 1995 through December 1997. From May 1995 through September 1996, webs 1 and 4 were considered controls. Captured mice from these webs were identified, marked, weighed, and measured, but not bled. Beginning in November 1996, we began collecting blood samples from mice on web 4. The bleeding process had little effect on survival (1). The methods for obtaining blood samples and the serologic testing of samples for hantavirus antibodies are described in Mills et al. (this issue, pp. 95-101). We examined population dynamics of common species infected with Sin Nombre virus (SNV) using data from three webs that were trapped continuously from May 1995 through December 1997. Using the minimum number of rodents known to be alive during a 3-day trapping session, we calculated an index of population size by taking the total number of rodents captured during each 3-day trapping session and adding to that sum the number of rodents captured on at least one previous and one subsequent session (2). The minimum number of hantavirus antibody–positive rodents was calculated in the same way. We estimated standing prevalence for each trapping session by dividing the minimum number of antibody-positive rodents by the minimum number of rodents known to be alive. Capture histories were used to estimate survivorship of the trappable population. These estimates were calculated as the percentage of rodents known to be alive a given number of months after initial capture. Although we refer to these estimates as survival rates, they are more accurately described as trapping web residency rates, as deaths cannot be distinguished from emigration. Trapping Results Between May 1995 and December 1997, 1,234 rodents were captured a total of 3,226 times, and 1,231 blood samples were obtained (Table 1). Bailey's pocket mouse (Chaetodipus baileyi) was the most common species captured (57% of rodents captured). Common murid rodents captured included white-throated wood rat (Neotoma albigula) (10%) and four species in the genus Peromyscus (27%). The cactus mouse (P. eremicus) was the most common Peromyscus species captured (12%) followed closely by the brush mouse (P. boylii) (11.5%). Deer mice (P. maniculatus) and white-footed mice (P. leucopus) were also captured but in low numbers (<3% each). Other species captured infrequently (<1%) included the fulvous harvest mouse (Reithrodontomys fulvescens), yellow-nosed cotton rat (Sigmodon ochrognathus), desert pocket mouse (C. penicillatus), and Merriam's kangaroo rat (Dipodomys merriami). Table 1. Prevalence of antibodies to Sin Nombre virus among wild rodents in southeastern Arizona, May 1995–December 1997 -------------------------------------------------------------------------- No. rodents trapped and released No. (total No. positive Family/Species Common name captures)(sup a) tested (%) -------------------------------------------------------------------------- Heteromyidae Dipodomys merriamiMerriam's 1 (2) 0 0 (0.0) kangaroo rat Chaetodipus spp. Pocket mice C. baileyi Bailey's pocket 704 (715) 329 0 (0.0) mouse C. penicillatus Desert pocket 25 (27) 7 0 (0.0) mouse Subtotal 730 (744) 336 0 (0.0) Muridae Neotoma albigula White-throated 126 (126) 51 0 (0.0) wood rat Onychomys torridusSouthern 7 (7) 7 0 (0.0) grasshopper mouse Peromyscus spp. White-footed mice P. boylii Brush mouse 137 (142) 98 13 (13.3) P. eremicus Cactus mouse 151 (152) 118 1 (0.8) P. leucopus White-footed 29 (30) 21 3 (14.3) mouse P. maniculatus Deer mouse 6 (6) 2 0 (0.0) Reithrodontomys Fulvous harvest fulvescens mouse 16 (16) 12 0 (0.0) Sigmodon Yellow-nosed 11 ochrognathus cotton rat (11) (11) 5 0 (0.0) Subtotal 483 (490) 314 17 (5.4) Total 1,213 (1,234) 650 17 (2.6) -------------------------------------------------------------------------- aTotal captures include rodents trapped and released and those that died during handling. Prevalence of Antibody-Positive Rodents Only rodents in the genus Peromyscus had antibodies reactive with SNV; however, antibody prevalence varied considerably among species within this genus (Table 1). Most (13 of 17) antibody-positive rodents were brush mice. One cactus mouse and three white-footed mice were also antibody positive. With the exception of one white-footed mouse, all antibody-positive rodents were captured in oak riparian vegetation. Antibody-positive rodents were captured on all three webs from which animals were bled; however, most (65%) were first captured on web 2 early in the study (May to June 1995). The farthest distance between trap stations where these web 2–rodents were captured was approximately 190 m, and half were captured at three adjacent trap stations along one transect line. All antibody-positive rodents were positive upon first capture, and most (58%) were never recaptured. Antibody-positive animals that were recaptured were caught an average of 3.8 times (standard deviation = 2.03, n = 7, range 2 to 8). All but one of the recaptured animals remained antibody positive on subsequent captures. The exception, a male brush mouse, was antibody negative on its three recaptures. Characteristics of Infected Populations Antibody-positive rodents were more likely to be male than female and were predominately adult (Table 2). The ratio of male to female among antibody-positive brush mice was significantly higher than that among the total sample (chi-square with Yates' correction = 7.97, degrees of freedom = 1, p = 0.005), and significantly more adults were antibody positive than would be expected from the distribution of age classes among the total sample (chi-square = 9.69, df = 2, p = 0.002). Although the sample size is too small for significance testing, these patterns hold for white-footed mice as well (Table 2). Table 2. Distribution of antibody-positive versus all brush mice, cactus mice, and white-footed mice, by sex and age ------------------------------------------------------------------------------ Brush mice Cactus mice White-footed mice -------------------------------------------------------------- No. (%) Total No. (%) Total No. (%) Total Characteristic positive no. (%) positive no. (%) positive no. (%) ------------------------------------------------------------------------------ Sex Male 12 (92) 51 (52) 0 59 (50) 3 (100) 10 (48) Female 1 ( 8) 47 (48) 1 (100) 59 (50) 0 11 (52) Age Juvenile 0 12 (12) 0 22 (18) 0 2 (10) Young adult 1 (8) 38 (40) 0 48 (41) 0 3 (14) Adult 12 (92) 48 (48) 1 (100) 48 (41) 3 (100) 16 (76) ------------------------------------------------------------------------------ Brush Mice Population Dynamics and Temporal Pattern of Infection The number of brush mice varied both by season and by year. The minimum number known to be alive was relatively high during the first 10 months of the study, May 1995 through March 1996 (Figure 1). The number of brush mice declined during the spring of 1996 and remained low until the fall, when the numbers increased but never reached the levels of the previous year. Captures for the next year followed a similar pattern with increased numbers during fall and winter (October through March), followed by a steady spring decline and summer low. The minimum number of brush mice known to be infected was highest during the initial part of our study [fig 1] (Figure 1). Eleven of the 13 Figure 1. Population trends of brush hantavirus antibody–positive mice mice, as determined by the minimum were first captured between May and number known to be alive, Santa Rita September 1995, gradually Experimental Range, southeastern disappearing from the population. By Arizona, May 1995–December 1997. October 1996, no animals were known to be infected on any of our trapping webs. One new antibody–positive brush mouse was captured in November 1996 [fig 2] and another in November 1997. Figure 2. Survivorship functions Similarly, the estimated standing (percentage of brush mice known to prevalence of hantavirus antibody be alive after initial capture) ranged from 40% in May 1995 to 0% in based on recapture data, Santa Rita both October 1996 and April through Experimental Range, southeastern October 1997 (mean= 8.25%). Arizona, May 1995–December 1997. Male and female brush mice showed similar rates of survivorship with an approximately 50% turnover rate around 2 months after initial capture (Figure 2). Hantavirus antibody–positive mice did not survive quite as long; the 50% turnover rate occurred approximately 1 month after initial capture. By 6 months after first capture, approximately 80% of all rodents had disappeared. A small percentage of brush mice continued to be captured for more than 1 year after tagging. Conclusions The overall prevalence of antibodies reactive with SNV antigen varied considerably among wild rodents captured in southeastern Arizona between May 1995 and December 1997, from 0% for Heteromyidae to 5.4% for Muridae. Low prevalence within the heteromyids has been commonly documented (3-5). Of mice, only Peromyscus were seropositive at our study site. The mean antibody prevalence of 7% for all Peromyscus was similar to the mean prevalence reported from Kansas (6) and Montana (7), although lower than that at many other sites in Arizona and New Mexico (3,4). The low hantavirus-antibody prevalence at our site may be related to its location in Sonoran Desert semigrassland and its relatively low rainfall; Mills et al. (4) found that prevalence of SNV was lowest at altitudinal and climatic extremes. The primary Peromyscus species with evidence of hantavirus infection at the Santa Rita Experimental Range was the brush mouse, recently shown to be an important carrier of SNV or an SNV–related virus throughout the southwestern United States (4). Even within a single species, overall prevalence of hantavirus antibodies has been reported to vary widely among different regions and habitats and in different seasons and years. In samples of deer mice from sites throughout the southwestern United States, Mills et al. (4) found antibody prevalence of 0% to 50%. Within states, overall prevalence in deer mice was 9.5% to 38.6% at 10 sampled sites in New Mexico (3) and 0% to 50% in 34 counties in California (5). Several studies have indicated, as does ours, that the presence and number of antibody-positive mice are not evenly distributed. Although Peromyscus were trapped in both vegetation types within our study site, all but one of the antibody-positive mice were trapped in oak riparian vegetation, and most were trapped in one portion of one web. Similarly, Mills et al. (4) captured antibody-positive deer mice in only 21 of 41 sites where deer mice were captured, and hantavirus antibody-positive brush mice in only 9 of 17 sites. Our results suggest that the prevalence of antibody-positive animals may be correlated with different habitats and provide additional evidence for focality of hantavirus in "reservoir" populations (4). While our sample sizes are too small to determine statistical significance, they suggest a correlation between population size and prevalence of hantavirus antibody. The number of antibody-positive animals was highest when the population was decreasing from an abundance of Peromyscus in the spring of 1995, the most recent peak. This finding is in contrast to local studies in the Channel Islands (8), Montana (7), and the regional study of Mills et al. (4), which found no relationship between antibody prevalence and density of deer mice. However, Childs et al. (3) found higher antibody prevalence in pinyon-juniper vegetation in 1993, when evidence suggests that rodent densities were unusually high (9). Additional data from our long-term study and other studies should help determine whether any relationship between density and antibody prevalence exists and if so, what the related temporal patterns are. Population sizes of rodents in the Sonoran Desert of southeast Arizona, as in other areas with climatic extremes, are highly variable. The number of P. boylii at Santa Rita Experimental Range was initially high but declined over the course of our study (perhaps because of changes in annual rainfall). To reproduce, many desert rodents require green vegetation (10), often not available in semidesert grasslands and xeroriparian areas. Total annual rainfall at Santa Rita Experimental Range was higher than normal in the 2 years before the start of our study. Since 1995, annual rainfall has been approximately 8 cm to 10 cm below the norm (unpub. data). Petryszyn (11) has linked high variability of Peromyscus populations in the Sonoran Desert with extreme fluctuation in winter rainfall. Others (12) have indicated local population expansion and retraction in response to wetter and drier conditions. Finally, our results are consistent with those of other studies that show a higher prevalence of infection (as indicated by antibody) in male and sexually mature rodents. However, we did not observe direct signs of aggressive encounters or fighting among infected males, as observed by Childs et al. (13) for hantaviral infection in Rattus norvegicus. Field studies of hantavirus infection and wild rodent populations provide a rare opportunity for public health officials, virologists, and ecologists to better understand the dynamics of rodent populations and the interactions between disease, humans, small mammals, habitat, and climatic factors. The few long-term datasets in ecology are invaluable for their contributions to the understanding of processes that vary in complex ways over time but are also relevant to management of both the natural environment and human health. Acknowledgments We thank T. Abeloe, C. Boal, M. Bucci, T. Cutler, L. Hall, C. Johnson, A. McLuckie, J. Martin, I. Rodden, and S. Simpson for assistance in the field. We also thank C. Levy, D. Engelthaler, J. Mills, T. Ksiazek, C. J. Peters, and J. Dunnum for assistance. R. Sanderson and C. Plumb at the Santa Rita Experimental Range provided logistical support. Funding for this study was provided by the Centers for Disease Control and Prevention and the Arizona Department of Health Services.